Marine Enzymes Biotechnology: Production and Industrial Applications, Part II - Marine Organisms Producing Enzymes [1st Edition] 9780128050798, 9780128047149

Table of contents :
Content:
Series PagePage ii
CopyrightPage iv
ContributorsPages ix-x
PrefacePage xiSe-Kwon Kim, Fidel Toldrá
Chapter One - Marine Microorganism: An Underexplored Source of l-AsparaginasePages 1-25A.A. Prihanto, M. Wakayama
Chapter Two - Marine Microbes as a Potential Source of Cellulolytic EnzymesPages 27-41N. Trivedi, C.R.K. Reddy, A.M. Lali
Chapter Three - Extremozymes from Marine ActinobacteriaPages 43-66J. Suriya, S. Bharathiraja, M. Krishnan, P. Manivasagan, S.-K. Kim
Chapter Four - Enzymes From Rare Actinobacterial StrainsPages 67-98J. Suriya, S. Bharathiraja, P. Manivasagan, S.-K. Kim
Chapter Five - Bioprospects of Microbial Enzymes from Mangrove-Associated Fungi and BacteriaPages 99-115K. Saravanakumar, N. Rajendran, K. Kathiresan, J. Chen
Chapter Six - Mechanism and Aquaculture Application of Teleost Enzymes Adapted at Low TemperaturePages 117-136C.-L. Wu, B.-Y. Li, J.-L. Wu, C.-F. Hui
Chapter Seven - Usefulness of Alginate Lyases Derived from Marine Organisms for the Preparation of Alginate Oligomers with Various BioactivitiesPages 137-160S. Takeshita, T. Oda
Chapter Eight - Marine Microbial Amylases: Properties and ApplicationsPages 161-177J. Suriya, S. Bharathiraja, M. Krishnan, P. Manivasagan, S.-K. Kim
Chapter Nine - Enzyme Immobilization: An Overview on Methods, Support Material, and Applications of Immobilized EnzymesPages 179-211V.L. Sirisha, Ankita Jain, Amita Jain
IndexPages 213-217

Citation preview

ADVISORY BOARDS KEN BUCKLE University of New South Wales, Australia

MARY ELLEN CAMIRE University of Maine, USA

ROGER CLEMENS University of Southern California, USA

HILDEGARDE HEYMANN University of California, Davis, USA

ROBERT HUTKINS University of Nebraska, USA

RONALD JACKSON Brock University, Canada

HUUB LELIEVELD Global Harmonization Initiative, The Netherlands

DARYL B. LUND University of Wisconsin, USA

CONNIE WEAVER Purdue University, USA

RONALD WROLSTAD Oregon State University, USA

SERIES EDITORS GEORGE F. STEWART

(1948–1982)

EMIL M. MRAK

(1948–1987)

C. O. CHICHESTER

(1959–1988)

BERNARD S. SCHWEIGERT (1984–1988) JOHN E. KINSELLA

(1989–1993)

STEVE L. TAYLOR

(1995–2011)

JEYAKUMAR HENRY

(2011–2016)

FIDEL TOLDRÁ

(2016– )

Academic Press is an imprint of Elsevier 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States 525 B Street, Suite 1800, San Diego, CA 92101-4495, United States The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom 125 London Wall, London, EC2Y 5AS, United Kingdom First edition 2016 Copyright © 2016 Elsevier Inc. 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. ISBN: 978-0-12-804714-9 ISSN: 1043-4526 For information on all Academic Press publications visit our website at https://www.elsevier.com/

Publisher: Zoe Kruze Acquisition Editor: Alex White Editorial Project Manager: Helene Kabes Production Project Manager: Surya Narayanan Jayachandran Cover Designer: Christian Bilbow Typeset by SPi Global, India

CONTRIBUTORS S. Bharathiraja CAS in Marine Biology, Annamalai University, Porto Novo, Tamil Nadu, India J. Chen State Key Laboratory of Microbial Metabolism, School of Agriculture and Biology, Shanghai Jiao Tong University, Shanghai, China C.-F. Hui Institute of Cellular and Organismic Biology, Academia Sinica, Taipei, Taiwan Amita Jain UM-DAE Centre for Excellence in Basic Sciences, University of Mumbai, Mumbai; D.Y. Patil University, Navi Mumbai, India Ankita Jain UM-DAE Centre for Excellence in Basic Sciences, University of Mumbai, Mumbai; University of Rajasthan, Jaipur, India K. Kathiresan CAS in Marine Biology, Faculty of Marine Sciences, Annamalai University, Parangipettai, India S.-K. Kim Marine Bioprocess Research Center; Specialized Graduate School Science & Technology Convergence, Pukyong National University, Busan, Republic of Korea M. Krishnan School of Environmental Sciences, Bharathidasan University, Tiruchirappalli, Tamil Nadu, India A.M. Lali DBT-ICT Centre for Energy Biosciences, Institute of Chemical Technology, Mumbai, India B.-Y. Li Institute of Cellular and Organismic Biology, Academia Sinica, Taipei, Taiwan P. Manivasagan Marine Bioprocess Research Center, Pukyong National University, Busan, Republic of Korea T. Oda Graduate School of Fisheries Science & Environmental Studies, Nagasaki University, Nagasaki, Japan A.A. Prihanto Faculty of Fisheries and Marine Science, Brawijaya University, Malang, Indonesia

ix

x

Contributors

N. Rajendran Government Arts College, Chidambaram, India C.R.K. Reddy Division of Marine Biotechnology and Ecology, CSIR-Central Salt and Marine Chemicals Research Institute, Bhavnagar; Academy of Scientific and Innovative Research (AcSIR), New Delhi, India K. Saravanakumar State Key Laboratory of Microbial Metabolism, School of Agriculture and Biology, Shanghai Jiao Tong University, Shanghai, China V.L. Sirisha UM-DAE Centre for Excellence in Basic Sciences, University of Mumbai, Mumbai, India J. Suriya School of Environmental Sciences, Bharathidasan University, Tiruchirappalli, Tamil Nadu, India S. Takeshita Center for Industry, University and Government Cooperation, Nagasaki University, Nagasaki, Japan N. Trivedi Division of Marine Biotechnology and Ecology, CSIR-Central Salt and Marine Chemicals Research Institute, Bhavnagar; Academy of Scientific and Innovative Research (AcSIR), New Delhi, India M. Wakayama College of Life Sciences, Ritsumeikan University, Shiga, Japan C.-L. Wu Institute of Cellular and Organismic Biology, Academia Sinica, Taipei, Taiwan J.-L. Wu Institute of Cellular and Organismic Biology, Academia Sinica, Taipei, Taiwan

PREFACE In the last decades, the progress on the knowledge of marine enzymes has advanced exponentially. This growing interest on marine enzymes is based on their relevant and unique properties that make them quite attractive because somehow they are different from the well-known terrestrial enzymes. Marine organisms may have to face extreme environmental conditions and this makes that most of their enzymes be active and stable under extreme conditions like very high or very low temperatures, high pressure, tolerance to high salt concentration, stability to acid or basic pH, easy adaptation to cold conditions, etc. All these properties make marine enzymes very attractive for new catalytic reactions and, of course, new applications in food and nutrition. In view of this increased interest, Advances in Food and Nutrition Research is publishing three consecutive volumes focused on the topic Marine Enzymes Biotechnology: Production and Industrial Application. Volume 78, which was recently published, corresponded to Part I that was mainly dealing with the production of enzymes from marine sources, this volume 79 corresponds to Part II dealing with the marine organisms producing enzymes, and volume 80 will correspond to Part III dealing with the applications of marine enzymes. This volume brings a variety of chapters reporting the marine organisms producing enzymes. So, this volume includes marine bacterial amylases, cellulases and alginate lyases, extremozymes from marine actinobacteria, enzymes from rare actinobacterial species, enzymes from mangrove-associated fungi and bacteria, and teleost enzymes adapted to low temperature. Methods for enzyme immobilization, types of supports, and applications are also discussed. This volume presents the combined effort of 26 professionals with diverse expertise and background. The Guest Editors wish to thank the publisher production staff and all the contributors for sharing their experience and for making this book possible. SE-KWON KIM FIDEL TOLDRA´ Guest Editors

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CHAPTER ONE

Marine Microorganism: An Underexplored Source of L-Asparaginase A.A. Prihanto*,1, M. Wakayama† *Faculty of Fisheries and Marine Science, Brawijaya University, Malang, Indonesia † College of Life Sciences, Ritsumeikan University, Shiga, Japan 1 Corresponding author: e-mail address: [email protected]

Contents 1. 2. 3. 4. 5. 6.

Introduction Short Historical View Occurrence and Distribution Structure and Properties of L-Asparaginase Production of L-Asparaginase Application of L-Asparaginase 6.1 Medicine 6.2 Food 7. Marine Environment as a Source of L-Asparaginase 7.1 Why Marine? 7.2 L-Asparaginase Derived from Marine Microorganisms 7.3 Challenge and Future Perspective of Marine Microorganism Exploration 8. Concluding Remarks References

2 2 3 4 8 11 11 12 13 14 15 16 20 21

Abstract L-Asparaginase

(EC 3.5.1.1) is an enzyme that catalyzes the hydrolysis of L-asparagine to acid. This enzyme has an important role in medicine and food. L-Asparaginase is a potential drug in cancer therapy. Furthermore, it is also applied for reducing acrylamide, a carcinogenic compound in baked and fried foods. Until now, approved L-asparaginases for both applications are few due to their lack of appropriate properties. As a result, researchers have been enthusiastically seeking new sources of enzyme with better performance. A great number of terrestrial L-asparaginase-producing microorganisms have been reported but unfortunately, almost all failed to meet criteria for cancer therapy and acrylamide reducing agent. As a largest area than Earth, marine environment, by contrast, has not been optimally explored yet. So far, a great challenge facing an exploration of marine microorganisms is mainly due to their harsh, mysterious, and dangerous environment. It is clear that marine environment, a gigantic potential source for marine natural products is scantily revealed, although several approaches L-aspartic

Advances in Food and Nutrition Research, Volume 79 ISSN 1043-4526 http://dx.doi.org/10.1016/bs.afnr.2016.07.005

#

2016 Elsevier Inc. All rights reserved.

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A.A. Prihanto and M. Wakayama

and technologies have been developed. This chapter presents the historical of L-asparaginase discovery and applications. It is also discussed, how the marine environment, even though offering a great potency but is still one of the less explored area for L-asparaginase-producing microorganisms.

1. INTRODUCTION L-Asparaginase

(EC 3.5.1.1) is an enzyme that catalyzes the hydrolysis of L-asparagine to L-aspartic acid. The intense exploration of the enzyme was firstly inspired by the anticancer activity of L-asparaginase. The application of L-asparaginase is not only to medicine but also recently used in foods because it is able to reduce acrylamide in several foods. However, due to its limitations, L-asparaginase has seldom been approved for use in medicine and food production. L-Asparaginases derived from Escherichia coli and Erwinia chrysanthemi have been produced and are commercially available. Application of L-asparaginase in cancer drugs is currently limited because of their lack of appropriate properties; specifically, short half-life, immune response development, allergic reaction, and provocation of anaphylactic shock. Furthermore, other L-asparaginases are usually accompanied by their high Km value. Therefore, the discovery of new L-asparaginases with better physiological and biochemical properties may result in a cancer cure and the elimination of acrylamide in food. The marine environment is a mega-biodiversity reservoir that covers >71% of the Earth with most of the area unexplored. This environment comprises various habitats, ranging from the surface to the bottom floor. Vast biodiversity in marine habitats is very surprising. It is estimated that the total number of prokaryotic cells in the oceans is 1029 (Whitman, Coleman, & Wiebe, 1998). Unfortunately, we know little about their potential. In this chapter, we present a comprehensive review of L-asparaginase along with its applications. The potential exploration of the marine environment for the possibility of obtaining L-asparaginase with new properties is also discussed.

2. SHORT HISTORICAL VIEW L-Asparaginase was detected for the first time in 1904 by Lang, who found asparagine-hydrolysis activity in cow tissue. Six years later, in 1910,

Marine Microorganism

3

Furth and Friedmann reported that horse and pig organs exhibited L-asparaginase activity, and they concluded that all animal tissues had the same level of L-asparaginase activity (Furth & Friedmann, 1910). In contrast, Clementi stated that the L-asparaginase activity was only exhibited in the specific organs of omnivores (in the case of pigs, it was in the liver), where carnivores, amphibians mammals, and reptiles were not included and only herbivores showed the L-asparaginase activity in all of their organs. Clementi also described a basic step for the application of this enzyme is anticancer therapy (Clementi, 1922). The application of L-asparaginase gained attention for the first time in 1922 when Clementi reported the previously unrealized-antileukemia activity of pig serum, which was corroborated by Kidd’s experiment which found that the pig-blood serum inhibited lymphosarcoma proliferation (Kidd, 1953). However, it was not until 1963 when researchers began to realize that L-asparaginase activity in pig serum was responsible for the serum’s cancer-inhibition activity (Broome, 1963). Since then, intensive efforts to discover an effective L-asparaginase for treating cancer have been conducted. It is not clear when L-asparaginase was introduced in food production. Friedman (2003) was the first researcher who suggested to use L-asparaginase for eliminating acrylamide in foods. However, clear evidence on the reduction ability of L-asparaginase toward acrylamide in food was reported by Novozyme researchers (Hendriksen, Kornbrust, Ernst, Stringer, & Heldt-Hansen, 2005). One year later, Ciesarova and coworkers reported the acrylamide mitigating effect of L-asparaginase (Ciesarova, Kiss, & Boegl, 2006).

3. OCCURRENCE AND DISTRIBUTION The widespread distribution of L-asparaginase has been reported. This enzyme can be found in almost all living organisms. Higher organisms, either animal or plants are known to be sources of L-asparaginase. Several higher organisms that exhibit L-asparaginase are presented in Table 1. Screening of 34 plants of the Solanaceae and Fabaceae families indicated that all exhibited L-asparaginase activity. L-Asparaginase is predominantly isolated from developing leaves and seeds. In the first development stage of plant cells, the product of the L-asparaginase reaction, aspartic acid is predictably involved in lysine, threonine, and methionine synthesis (Bruneau, Chapman, & Marsolais, 2006).

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A.A. Prihanto and M. Wakayama

Table 1 Occurrence of L-Asparaginase in Higher Organisms High Organism Organism References

Animal

Guinea-pig

Rogez, Plaquet, and Biserte (1975)

Plant

Erwinia carotovora

Howard and Carpenter (1972)

Lupinus arboreus

Lough et al. (1992)

Lupinus angustifolius

Dickson et al. (1992)

Pisum sativum L.

Chagas and Sodek (2001)

Arabidobsis taliana

Casado et al. (1995)

Capsicum annum L. Bano and Sivaramakrishnan (1980) Tamarindus indica

Bano and Sivaramakrishnan (1980)

Soybean

Tonin and Sodek (1990)

Lotus japonicus

Credali et al. (2011)

Withania somnifera Oza, Trivedi, Parma, and Subramanian (2009)

Microorganisms are considered to be the most abundant sources of In addition, the large-scale production of L-asparaginase is not very cumbersome because microorganisms are easily cultured and can rapidly grow. Moreover, the downstream process of enzymes derived from microorganism is relatively straightforward. L-asparaginase.

4. STRUCTURE AND PROPERTIES OF L-ASPARAGINASE On the basis of amino acid sequence analysis, L-asparaginase has been categorized into three types: the bacterial, plant, and Rhizobacterium etli type. The bacterial type is less homologous compared with plant type. No homology with the other types was found in R. etli-asparaginase derived from the AnsA gene, and it has very low similarity with other known bacterial L-asparaginases. In this chapter, we discuss only the bacterial asparaginase I (AnsI) and asparaginase II (AnsII) and plant-type asparaginases. L-Asparaginases can be classified into two types (Table 2). AnsII is more studied than other types L-asparaginases due to their superior application in cancer. Hence, for example, crystallographic data for AnsI from Pyrococcus horikoshii was firsly available in 2005 (Yao, Yasutake, Morita, & Tanaka, 2005) while AnsII from Acinetobacter glutaminasificans was earlier (Tanaka et al., 1988).

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Marine Microorganism

Table 2 Properties of Bacterial Type-L-Asparaginases Properties AnsI

AnsII

Location

Cytoplasmic

Periplasmic

Affinity

Low (3.5 mM)

High (15 μM)

Protein form

Dimer, tetramer

Tetramer

Half-life in serum

Short (15 min)

Long (8–49 h)a

a Route of administration-dependents, intravenous administration gave shorter half-life than that intramuscular.

Various L-asparaginase genes have been identified, cloned, and sequenced. Multiple sequence analysis of their genes showed relatively low similarity (Fig. 1). The highest preserved amino acids were found between AnsII of bacterial asparaginases, E. coli, plant-type asparaginase, and E. chrysanthemi. The similarity between E. coli AsnII and Erwinia asparaginase is 48%, and the similarity for E. coli AsnI is only 21%. Low similarities between asparaginase from R. etli and other asparaginases (7–9%) have been observed. The three-dimensional structures of both AnsI and AnsII have been determined. The two most successful and famous L-asparaginases, AsnII derived from E. coli (EcA) and E. crysanthemi (ErA) were used for X-ray crystallographic analysis (Fig. 2). EcA is a homotetramer with about 330 amino acids in each subunit. The N-terminal domain of the subunit contains 4 α-helices and 10 β-sheets. Nβ7 and Nβ8 form an antiparallel β hairpin by leaving the main N-terminal sheet. On the other hand, the C-terminal contains the same amounts of α and β forms. This hairpin has been deduced to have an important role in subunit adhesion. ˚. The crystal structure of EcA has been determined at a resolution of 2.3 A In all AnsII enzymes, the access to the active site is covered by a flexible loop. The surface loop will be open to allow the substrate to bind, after which the active site loop is closed (Aung, Bocol, Schleper, & R€ ohm, 2000). On the basis of crystallographic data, the active site was determined to be located between the N-terminal of one site and the C-terminal of other sites. Even though the threonine residue in the position of 12 (Thr12) is not involved in the substrate binding process, Thr12 and threonine 89 (Thr89) were the nucleophilic sites (Harms, Wehner, Aung, & R€ ohm, 1991). Thr198 might be involved in formation of a hydrogen bond that stabilizes and links the Nand C-terminal domains. The unique structure of EcA is the left-handed crossover form. The left-handed crossover in the N-terminal β-sheet is considered to be rare in protein structures.

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A.A. Prihanto and M. Wakayama

Fig. 1 Multiple alignment sequences of L-asparaginase classes.

Fig. 2 Structure of L-asparaginase. Ribbon structure of AnsII from Escherichia coli, PDB ID: 3ECA (A). L-Asparaginase II (periplasmic) from Erwinia crhysanthemi, PDB ID: 1JSR (B).

Marine Microorganism

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Two different subtypes of plant asparaginases according to their dependence on K+ for catalytic activity have been reported. These two categories, namely K+-dependent and K+-independent asparaginases are classified according to their roles in asparagine catabolism of plants. One role is deamidation, which mainly occurs in developing plant organs, such as roots and leaves, and the other is transamination, which mainly occurs in mature plants. Even though the subtypes have different classifications, both share similar nucleotides sequences. High homology was found between K+independent L-asparaginases from Lupinus luteus (L1A) and bacterial, E. coli L-asparaginase (EcIII). Ribbon structure of L1A is depicted in Fig. 3. The plant L-asparaginases, L1A and EcIII, belong to the superfamily of Ntn hydrolase. Both L-asparaginases are produced in inactive forms, which require an autocatalytic process to become active enzymes. A comprehensive review for this autocatalytic process was conducted by Michalska and Jaskolski (2006). As members of the Ntn-hydrolase family, these L-asparaginases have a classical form, αβ heterodimer with a sandwich-like (αββα) form. Therefore, these L-asparaginases are tetramers with each subunit containing α (residue 1–192) and β (residue 193–325) units. Threonine has been shown to have an important role in the catalytic process. Thr139 and Thr179 are a nucleophilic residue in L1A and EcIII, respectively.

Fig. 3 Structure of plant-type asparaginase. Ribbon structure of L1A from Lupinus luteus PDB ID: 2GEZ.

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5. PRODUCTION OF L-ASPARAGINASE The production improvement has come from different approaches, from simple to very complicated. Fermentation processes can be categorized into solid-state fermentation (SSF) and submerged fermentation (SF). SSF is usually used for metabolite production. SSF offers more advantages than does SF. SSF requires less control process, low energy with negligible aeration, relatively shortcoming downstream process; hence, the operation is easier and simpler. Furthermore, SSF also generates less effluent, which minimizes waste and pollution. However, the substrate should be in a sufficiently moist form in SFF. This requirement will eventually limit the application of SSF only to fungi and microorganisms which require lowmoisture media. Therefore, this system is not particularly suitable for bacteria which usually require high water activity (aw). The most common approach to increase enzyme production is modification of the culture media. The interaction between medium components and culture environments is species dependent and will require further exploration to attain the best conditions for enzyme production. The process parameters that commonly have an impact on the production of enzymes are nitrogen and carbon sources, temperature of incubation, pH of the medium, culture time, aeration, and/or agitation if necessary and percentage of additional inoculum (v/v) into the main culture. SSF can use agricultural wastes or other underutilized materials for medium. These approaches are usually developed by design of experiment using statistical methods. This idea is interesting because this venture uses “wastes or underutilized materials” (WUM) to support the production of more valuable materials. The reports are summarized in Table 3. In the utilization of WUM for fermentation media, the nutritional requirements and characteristics of the microorganisms should be the main consideration. If the WUM cannot be directly metabolized, pretreatments might be necessary to enable easy metabolism by microorganisms. WUM is mainly used as carbon source and it is rare to use WUM as nitrogen source. The availability of carbon sources in industrial and agricultural wastes is greater than that of nitrogen sources. Growth culture modification based on a deep understanding of a microorganism’s metabolism system and/or gene recombinant has been introduced. One example is culturing of bacteria with an oxygen vector. Involvement of an oxygen vector in the production of L-asparaginase from

Table 3 Optimization of L-Asparaginase Production Using SSF No Species WUM Fermentation Conditions

Production

References

1

Cladosporium sp. Wheat bran, rice 120 h fermentation time under aerobic 3.74 U bran, and bagasse condition, moisture content of 58%, pH 5.8, incubation temperature of 30°C

Kumar, Ramasamy, and Manonmani (2013)

2

Escherichia coli ATCC 11303

Lactose

Kenari, Alemzadeh, and Maghsodi (2011)

3

Bacillus licheniformis (RAM-8)

–

4

Pseudomonas aeruginosa

Casein and corn steep liquor

pH 7.9; casein hydrolysate, 3.11%; and corn steep liquor, 3.68%

5

Bacillus cereus MAB5

Soyabean meal and wood chips

Soyabean meal (6.282 8 g/L), aspargine 51.54 IU/mL (5.5 g/L), wood chips (1.383 8 g/L), and NaCl (4.535 4 g/L)

6

Serratia marcescens (NCIM 2919)

Coconut oil cake Coconut oil cake 7.6 g/L, moisture content 50%, temperature 35.5°C, and pH 7.4

7

Aspergillus niger

Leguminous crops

0.08% lactose, 1.79% tryptone, 1.6% yeast 1.03 U/mL extract, 2% KH2PO4, and 0.19% L-asparagine 29.94 IU/ml

Mahajan, Saran, Kameswaran, Kumar, and Saxena (2012)

142.8 IU

Abdel-Fattah and Olaman (2002)

5.86 U/gds

Thenmozhi, Sankar, Karuppiah, and Sampathkumar (2011) Ghosh, Murthy, Govindasamy, and Chandrasekaran (2013)

A 96-h fermentation time under aerobic, 0.9  3.35 U/g Misra (2006) of dry substrate moisture content of 70%, 30 min of cooking time and 1205–1405 μ range of particle size, pH 6.5, and temperature 40°C Continued

Table 3 Optimization of L-Asparaginase Production Using SSF—cont’d No Species WUM Fermentation Conditions

8

Aspergillus wentii Palm oil cake

9

Bacillus subtilis

A 48 h fermentation time under aerobic, 89.954 IU moisture content of 89.51%, inoculum volume 30%, temperature 34°C, and pH 5.8

Ground nut cake 48 h of incubation at temperature 37°C moisture content of 70% and pH 7

10 Fusarium equiseti Soybean meal

Production

18.4 U/ml

8.51 IU Incubation period (48 h), moisture content (70%, v/w), particle size (3 mm), inoculum volume (20%, v/w), supplemented with glucose (0.5%, w/v), ammonium sulfate (0.5%, w/v), yeast extract (0.5%, w/v), and temperature 28°C

References

Satya, Anuradha, and Reddy (2014)

Shukla and Mandal (2012) Hosamani and Kaliwal (2011)

Marine Microorganism

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Bacillus brevis enhances the production of the enzyme. Liquid paraffin at a concentration of 4% enhances the production of the enzyme and cell mass by 34% and 48%, respectively (Narta, Roy, Kanwar, & Azmi, 2011). The role of an oxygen vector in the fermentation is to maintain an adequate concentration of dissolved oxygen. Manipulation of culture conditions by stress and shock stimulation can be used to enhance the production of L-asparaginase. In fact, significant enhancement of L-asparaginase production in Arthrospira platensis under NaCl and temperature shock culture conditions without UV light has been described. This condition initiates the requirement of energy generation by degrading the nitrogen stock, cyanophycin, which causes an increase in ASNase production (Prihanto & Wakayama, 2014). Medium modification, by changing glucose to maltose will increase production of L-asparaginase in E. coli. This modification is based on the fact that glucose acts as an inhibitor of AnsB gene expression that is regulated by the catabolite activator protein, which is also called cyclic AMP (cAMP) receptor protein. A high level of cAMP due to a high level of glucose will somehow hamper AnsB expression. The disaccharide, maltose, is used for bacterial growth and does not interfere with expression of AnsB (Ghasemi et al., 2008). Researchers have also found that elimination of Na2HPO4 in M9 medium improved productivity of L-asparaginase in bacteria by reducing the buffer properties and eliminating the bias of ammonia. Advanced technology for supporting the production of L-asparaginase has also been reported. These techniques are mainly performed after an enzyme has shown superior properties. Cloning of the Thermococcus kodakaraensis gene (TK1656) encoding L-asparaginase resulted in L-asparaginase with good properties. This enzyme exhibits high activity at a temperature of 85°C, active toward D-asparagine, and possesses high thermostability (Chohan & Rashid, 2013).

6. APPLICATION OF L-ASPARAGINASE 6.1 Medicine Cancer cells cannot synthesize asparagine although in normal cells asparagine can be produced inside the cell by asparagine synthase. To meet the requirements of cells for asparagine, cancer cells rely on asparagine supplied directly from blood. By limiting the supply of asparagine, the growth of the cancer cells is inhibited. Hence, cell proliferation can be stopped. In vitro studies have shown that L-asparaginase inhibited several different cancer cell types

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(i.e., acute lympoblastic leukemia, lymposarcoma, melanosarcoma, and NK-cell lymphoma) (Avramis, 2011; Yong, Zheng, Zhnag, & Zhu, 2003). Even with these promising in vitro results and the abundant sources reported for asparaginase, only two sources of L-asparaginase are commercially available. The only approved L-asparaginases for cancer treatment are ones derived from E. coli and E. chrysanthemi. Several brands of these enzymes are available in the market; for example, ELSPAR, ONCASPAR, ERWINAZE, and the less popular, KIDROLASE. All of these commercial drugs are not without disadvantages. L-Asparaginase drugs show several side effects due to unsuitable properties when administered for long periods (mostly >25 weeks). Administration of L-asparaginase to the cancer patients has been reported to cause acute pancreatitis (Chiewchengchol, Wanankul, & Noppakun, 2009). Furthermore, increases in cholesterol levels in acute lymphoblastic leukemia patients receiving L-asparaginase treatment have been also reported (Cohen, Bielorai, Harats, Toren, & Hamiel, 2010). L-Asparaginase from E. coli causes allergic reactions and forms antiasparaginase antibodies (Zalewska-Szewczyk, Andrzejewski, & Bodalski, 2004). L-Asparaginase from Erwinia sp. is less allergenic and free from antibody problems, but exhibits a shorter half-life in serum as a result of proteolysis activity by trypsin through cleavage of the peptide bond between Lysn53–Gly54 and Arg206–Ser207 in C-terminal region (Kotzia, Lappa, & Labrou, 2007). Furthermore, the intrinsic glutaminase activity of Erwinia L-asparaginase increases its toxicity (Ramya, Doble, Rekha, & Pulicherla, 2012; Warrell et al., 1982). To overcome this situation, an effort is the development of Lasparaginase conjugated to polyethylene glycol, called PEGASPARGASE, and has a longer half-life. It is more stable at high temperature and over a wide pH range; however, it still causes the anaphylactic reactions (Sahiner et al., 2013). Indeed, L-asparaginase is promising in the management of cancer even though needs improvement in view of all those drawbacks.

6.2 Food After discovery of acrylamide in food by Tareke, a Swedish researcher in 2002, acrylamide in foods became a global concern, mainly because of its carcinogenic effect. Food scientists have been focused to reduce and mitigate the formation of acrylamide in foods. Several approaches and processes have been designed for example, avoiding the involvement of acrylamide

Marine Microorganism

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precursors in the process by selecting raw materials with low levels of asparagine, soaking, storage under low temperature, and blanching. Enzymatic hydrolysis of L-asparagine to L-aspartic acid by L-asparaginase or other amidases may result in low-acrylamide content in foods. The first practice to reduce acrylamide using asparaginase was initiated by Novozymes researchers in 2005. They found that application of L-asparaginase from Aspergillus oryzae to French fries, biscuits, crisp bread, and fabricated chips, efficiently reduced formation of acrylamide (Hendriksen et al., 2005). Dough pastry, which is treated using asparaginase, is able to convert 96–97% of L-asparagine to L-aspartic acid, and as a consequence, the level of formed acrylamide can be reduced up to 90% (Kukurova, Morales, Bedna´rikova, & Ciesarova, 2009). Combination of physical and enzymatic methods has also been applied. The application of L-asparaginase from Bacillus subtilis along with the pretreatments, decreased formation of acrylamide by 80% in sliced potatoes (Onishi et al., 2015). Even more, 90–97% reduction of acrylamide in foods made from dried potato powder has been achieved (Ciesarova et al., 2006). The form and texture of raw materials significantly affect acrylamide formation because of the penetration factor. Unfortunately, efficiency of L-asparaginase for acrylamide reduction agents has mostly been achieved in only lab-scale experiments. In fact, these treatments sometimes give a different result in real food processing (largescale treatment), and lower efficiency has been usually reported. Commercial food grade L-asparaginases have been available since 2008. These have been obtained from Aspergillus niger under the name of DSM’s PreventASe® and A. oryzae (Novozyme’s Acrylaway). Both have been approved in several countries. At present, these are the only two products. Hence, further research is required to obtain new food grade L-asparaginases with superior properties.

7. MARINE ENVIRONMENT AS A SOURCE OF L-ASPARAGINASE For a long time, marine microorganisms have been overlooked and neglected. For many years, microorganisms from terrestrials are a dominant source for isolating and studying natural products. The straightforward screening, culture, and collection process of terrestrials’ microorganisms provoke scientists to focus only on terrestrial areas. The minor researches related to marine microorganisms are most probably due to the limitation

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methods to screen and isolate the marine microorganisms and the geophysical barriers. Unlike terrestrial’s microorganisms, there is a fact that most of marine microorganisms cannot be cultured. Furthermore, ocean is somewhat different in terms of their physical conditions, temperature, pH, hydrostatic pressure, etc. Hence, marine microorganisms are difficult but not impossible to be studied and explored.

7.1 Why Marine? The potential of marine microorganism is huge. The ocean covers one-third area of our planet. About 71% of earth surfaces with 3.6  188 km2 coup the area of Earth. The area covered by seawater is called ocean basin. The depth of ocean is ranging from the level of terrestrial coasts cover until 11 km depth found in ocean trenches. The deepest ocean basin, Mariana Trench was recorded in Pacific Ocean. Even with cutting edge technology in marine–vehicle engineering, this area is still difficult to be traveled, because of extreme pressure. It is one of many factors which contribute to the fact that most of marine environment, and their organisms, especially marine microorganisms still remain underexplored and mysterious. Whereas, marine enzymes offer unique and superior properties such as high salt tolerance, hyperthermostability, cold adaption, barophilic, etc. (Debashish, Malay, Barindra, & Joydeep, 2005). The ocean areas are divided into several categories. The illustration of ocean basin was depicted in Fig. 4. Based on topographical feature, the area of ocean is differentiated as follows: litoral, continental shelf, continental slope, abyssal plain, oceanic trench, volcanic ocean, and ocean ridges. Based on the depth of water, they can be divided into epipelagic, mesopelagic, bathypelagic, abyssopelagic zone and the deepest area in the ocean, hadal zone. Based on the penetration of the sunlight, three different zones are found: photic, mid-water, and aphotic zone. Each zone experienced the different biogeochemical processes, hence, each zone has its unique ecosystem directly affecting to the life of occupied marine organisms. Marine organisms display unique live characters due to their unique habitat. The fact that they experienced harsh environmental factors, and success adaptation to the oceanic process and its biogeochemical such as temp, oxygen cycling, carbon cycling, pressure, and pH due to the ocean acidification, etc., lead us to the prediction that they will develop and produce unique system and natural bioactive products. This adaptation process will result in the generation of natural products such as primary and secondary metabolites (Imhoff, Labes, & Wiese, 2011; Montaser & Luesch, 2011). All of

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Fig. 4 Cross-section profiles of ocean basin.

these make the ocean a tremendous biological active compounds reservoir. This expectation triggers many institutes, researchers, and even industrial managers around the world to boost the exploration of the ocean. Intensively explored area is only above the 200 m below surface (photic zone). The exploration in this zone is rather straightforward due to the little effect of physical barriers. This area is still light penetrable; the temperature is relatively warm and the pressure is not so high. Ocean pressure is the most difficult hindrance because every 10 m below the seawater surface, the pressure increases by one atmosphere (1 atm). More than 20 atm hydrostatic pressures will hit every living organism in the area below the 200 m depth. This is far beyond the human capability. To overcome this difficulties, the oceanographers, rely on the development of the submarine vessel that is able to penetrate the oceanic water columns and, of course, this exploration will be very costly. This suggests us that the ocean zones below the 200 m still hold enormous potential which is underexplored yet.

7.2 L-Asparaginase Derived from Marine Microorganisms A couple of years ago, explorations of marine organisms were usually focused on macroorganisms. The objective was to identify natural products

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that had potential as drugs. Later, researchers investigated the associated enzymes after recognition of the potential of the marine microorganisms for producing enzymes with unique and superior properties. For example, a study on polyketides synthase from marine microorganisms associated with marine sponges was driven by the drug potential of polyketides, which had been previously determined in marine sponges (Schirmer et al., 2005). From several studies, soil and marine seawater have been shown to be potential sources for L-asparaginase-producing marine bacteria. These microorganisms are abundant in marine soil. Recently, there has been a growth in the number of reported L-asparaginase-producing bacteria; however, they are considerably fewer than their counterparts, L-asparaginase-producing terrestrial microorganisms. The known L-asparaginase-producing marine-related microorganisms are listed in Table 4. Marine bacteria, archaea, and fungi have been reported to produce L-asparaginase and have been mostly isolated only from the most accessible parts of the ocean basin. They are from soil, including seaweeds, corals, and fish in intertidal and epipelagic zones. Only several archaea have been isolated from below the epipelagic zone. To date, the only reported L-asparaginase-producing bacterium isolated from the zone below the epipelagic zone is Bacillus aryabhattai. Detailed studies on L-asparaginase derived from marine microorganisms have not yet been reported.

7.3 Challenge and Future Perspective of Marine Microorganism Exploration While ongoing exploration of the marine microorganism in the epipelagic zones is still under intensive effort, the other research approaches show up. The exploration of the ocean basin was not only in the epipelagic zone but also the area far below the surface water. Marine microorganism offers unique and novel biochemical properties. For example, deep sea marine microorganisms show alkaliphiles, thermophiles, psychrophilic, and halophiles. Indeed, all of those extreme bacteria have a potential to produce extremozymes. However, the exploration of this area is difficult because of physical barriers such as temperature, the absence of light, and the hydrostatic pressure. The temperature in the deepest hadal zone is slightly above the freezing degree, about 4°C. The light was unable to penetrate after they reach the maximum depth at around 200–500 m. In the Mariana Trench which is considered as the deepest ocean basin, the pressure is above 1000 atm. Hence, exploring ocean is problematic, dangerous, and expensive.

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Marine Microorganism

Table 4 Marine-Related Microorganisms Which Were Reported to Produce L-Asparaginase in the Last Decade Bacteria Source Reference

Streptomyces sp.

Sediment

Sivasankar et al. (2013)

Streptomyces radiopugnans MS1

Sediment

Kumar and Selvam (2011)

Streptomyces sp. PDK7

Sediment

Dhevagi and Poorani (2006)

Streptomycete strain WS3/1

Sediment and coral

Kumari, Sankar, and Prabhakar (2011)

S. nousei MTCC 10469

Sponge

Dharmaraj (2011)

S. aureofasciculus

Estuarine-fish

Sahu, Sivakumar, Poorani, Thangaradjou, and Kannan (2007)

S. chattanoogenesis

Estuarine-fish

Sahu et al. (2007)

S. hawaiiensis

Estuarine-fish

Sahu et al. (2007)

S. orientalis

Estuarine-fish

Sahu et al. (2007)

S. canus

Estuarine-fish

Sahu et al. (2007)

S. olivoviridis

Estuarine-fish

Sahu et al. (2007)

Pseudomonas sp.

Sediment

Izadpanah Qeshmi, Javadpour, Malekzadeh, Jahromi, and Rahimzadeh (2014)

Bacillus sp.

Sediment

Izadpanah Qeshmi et al. (2014)

Zobellella sp.

Sediment

Izadpanah Qeshmi et al. (2014)

Pseudomonas sp.

Sediment

Izadpanah Qeshmi et al. (2014)

Oceanimonas sp.

Sediment

Izadpanah Qeshmi et al. (2014)

Pseudomonas sp.

Soil

Audipudi, Pallavi, and Supriya (2013)

Bacillus sp.

Soil

Audipudi et al. (2013)

Escherichia coli

Sediment

Sudha (2009)

Proteus sp.

Sediment

Sudha (2009)

Proteus vulgaris

Sediment

Sudha (2009)

Bacillus sp.

Sediment

Sudha (2009)

Pseudomonas sp.

Sediment

Sudha (2009) Continued

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Table 4 Marine-Related Microorganisms Which Were Reported to Produce L-Asparaginase in the Last Decade—cont’d Bacteria Source Reference Archaea

Pyrococcus furiosus

Sediment

Bansal, Gnaneswari, Mishra, and Kundu (2010)

Thermococcus kodakaraensis

Sediment and water

Chohan and Rashid (2013)

Pyrococcus horikoshii

Sediment

Yao et al. (2005)

Fusarium sp.

Seaweed

Thirunavukkarasu, Suryanarayanan, Murali, Ravishankar, and Gummadi (2011)

Alternaria sp.

Seaweed

Thirunavukkarasu et al. (2011)

Aspergillus sp.

Seaweed

Thirunavukkarasu et al. (2011)

Chaetomium sp.

Seaweed

Thirunavukkarasu et al. (2011)

Cladosporium sp.

Seaweed

Thirunavukkarasu et al. (2011)

Curvularia tuberculata

Seaweed

Thirunavukkarasu et al. (2011)

Drechslera sp.

Seaweed

Thirunavukkarasu et al. (2011)

Emericella nidulans

Seaweed

Thirunavukkarasu et al. (2011)

Nigrspora sp.

Seaweed

Thirunavukkarasu et al. (2011)

Penicillium sp.

Seaweed

Thirunavukkarasu et al. (2011)

Phoma sp.

Seaweed

Thirunavukkarasu et al. (2011)

Pestalotiopsis sp.

Seaweed

Thirunavukkarasu et al. (2011)

Pithomyces sp.

Seaweed

Thirunavukkarasu et al. (2011)

Aspergillus terreus

Sediment

Sudha (2009)

Aspergillus niger

Sediment

Sudha (2009)

Aspergillus fumigatus

Sediment

Sudha (2009)

Aspergillus luchuensis

Sediment

Sudha (2009)

Penicillium janthinellum

Sediment

Sudha (2009)

Penicillium citrinum

Sediment

Sudha (2009)

Curvularia palase

Sediment

Sudha (2009)

Fungi

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Marine Microorganism

Table 4 Marine-Related Microorganisms Which Were Reported to Produce L-Asparaginase in the Last Decade—cont’d Bacteria Source Reference

Trichoderma reesei

Sediment

Sudha (2009)

Trichoderma koningii

Sediment

Sudha (2009)

Trichoderma harzianum

Sediment

Sudha (2009)

Trichoderma virens

Sediment

Sudha (2009)

Trichoderma viridae

Sediment

Sudha (2009)

Trichoderma hamatum

Sediment

Sudha (2009)

Fusarium oxisporum

Sediment

Sudha (2009)

Drecheriella ellssi

Sediment

Sudha (2009)

Aspergillus sp.

Soil, mangrove Gupta, Dash, and Basak (2009)

Penicillum sp.

Soil, mangrove Gupta et al. (2009)

Fusarium sp.

Soil, mangrove Gupta et al. (2009)

Helminthosporium sp.

Soil, mangrove Gupta et al. (2009)

Scophulariopsis sp.

Soil, mangrove Gupta et al. (2009)

Paecilomyces sp.

Soil, mangrove Gupta et al. (2009)

Pestalotiopsis sp.

Soil, mangrove Gupta et al. (2009)

Several technologies were backing the exploration of marine microorganisms. Sending human to deep sea is impossible. The idea to build a machine to dive into a deep sea is costly. A robotic technology, remotely operated vehicles (ROVs) and autonomous underwater vehicles (AUVs) have answered these challenges. Some researchers have built ROVs which are cheaper, and more efficient. Several examples are “Seagliders” small ROV, reusable, and autonomous (Eriksen et al., 2001). Seabed AUV is able to travel up to 1000–2000 m (Sing, Can, Eustice, Lerner, & Mcphee, 2004). Later, Nereus, a hybrid robot, which is able to travel to Mariana Trench, was introduced in 2007 (Bowen et al., 2009). Unfortunately, in 2014, this vehicle was lost under the sea due to hydrostatic pressure. The laboratory culture of marine microbes is one of the big challenges. This difficulty is caused by the lack of technology to mimicking their natural habitat including physical and biochemical features. Therefore, if we are able to grow the microorganisms in the plate, they are only a few representative

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of the whole microbial biodiversity. This phenomena is called “The great plate count anomaly”. This term describes the lack of cultivable marine bacteria in a plate medium, and it usually represents 1 per 100 cells (Epstein, 2013; Staley & Konopka, 1985). From this phenomenon, we knew that culturing marine bacteria under standard medium and conventional culture method will produce only 1% of success. Several efforts offer the solution to this problem, such as medium and culture conditions improvement. Medium modification by changing the ionic ratio of sodium and potassium in the medium by nearly 1:1 is the best condition to culture marine bacteria isolated from fish (Simidu & Hasuo, 1968). The new technique by encapsulating single bacterial cells in microdroplets of solidified agarose under low nutrient flux condition was introduced by Zengler et al. (2002). They claimed that the percentage of microbial recovery of this method was nearly same with 16 ribosomal RNA results, ranging from 84.9% to 98.6% similarity, depending on the samples. Bollmann’s research introduced the use of diffusion chamber in order to increase the diversity of recovered microorganisms. These methods slightly bring a natural environment where the microorganisms live, in the laboratory, and mimicking their natural condition (Bollmann, Lewis, & Epstein, 2007). The majority of marine microorganisms remain uncultured. Molecularbased approach was finally taking their role to overcome the lack of microorganism exploration due to the culture techniques limitation. This method allows researchers to analyze all cells without growing them in the plate. This technique also can capture almost all the diversity of microorganisms. Although it still in early stage, molecular-based approach, such as proteomic, has shown it’s prospective to explore genes and its functional gene expressions. By using metagenomic analysis revealed that 6.12 million proteins from 7.7 million DNA sequencing reads (6.3 billion bp) data came from only marine planktonic microbiota (Yooseph et al., 2007). Unfortunately, all of these methods require the cutting edge technologies and they are very expensive so that all researchers might not have such requirements. Hence, in the future, the development of new methods, which offers simplicity and cost friendly with high reliability, should be taken into account.

8. CONCLUDING REMARKS Marine microorganisms are considered to be the most diverse organisms in the ocean basin. These microorganisms inhabit all seawater habitats

Marine Microorganism

21

of the ocean basin. Surprisingly, their potential has not been maximally explored. Rapid developments technologies, such as molecular biology may greatly contribute to successful ocean basin exploration to obtain L-asparaginase derived from marine microorganisms. There is no doubt that marine microorganisms are the next promising source for L-asparaginase production; however, marine exploration is hampered by various impediments. Over time, technological developments should lead to discovery of myriad novel marine L-asparaginases that have unique and superior properties.

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Prihanto, A. A., & Wakayama, M. (2014). Combination of environmental stress and localization of L-asparaginase in Arthrospira platensis for production improvement. 3 Biotech, 4, 647–653. Ramya, L. N., Doble, M., Rekha, V. P., & Pulicherla, K. (2012). L-Asparaginase as potent anti-leukemic agent and its significance of having reduced glutaminase side activity for better treatment of acute lymphoblastic leukaemia. Applied Biochemistry and Biotechnology, 167, 2144–2159. Rogez, J. C., Plaquet, R., & Biserte, G. (1975). Guine pig liver L-asparaginase. Separation, purification, and intracellular localisation of two distinct enzymatic activities. Biochimica et Biophysica Acta, 410, 370–381. Sahiner, U. M., Yavuz, S. T., G€ okce, M., Buyuktiryaki, B., Altan, I., Aytac, S., et al. (2013). Anaphylactic reaction to polyethylene-glycol conjugated-asparaginase: Premedication and desensitization may not be sufficient. Pediatrics International, 55, 531–533. Sahu, M. K., Sivakumar, K., Poorani, E., Thangaradjou, T., & Kannan, L. (2007). Studies on L-asparaginase enzyme of actinomycetes isolated from estuarine fishes. Journal of Environmental Biology, 28, 465–474. Satya, C. V., Anuradha, C., & Reddy, D. S. R. (2014). Palm oil cake: A potential substrate for L-asparaginase production. International Journal of Innovative Research in Science, Engineering and Technology, 3, 14627–14632. Schirmer, A., Gadkari, R., Reeves, C. D., Ibrahim, F., DeLong, E. F., & Hutchinson, C. R. (2005). Metagenomic analysis reveals diverse polyketide synthase gene clusters in microorganisms associated with the marine sponge Discodermia dissoluta. Applied and Environmental Microbiology, 71, 4840–4849. Shukla, S., & Mandal, S. K. (2012). production optimization of extracellular L-asparaginase through solid-state fermentation by isolated Bacillus subtilis. International Journal of Applied Biology and Pharmaceutical Technology, 4, 219–226. Simidu, U., & Hasuo, K. (1968). An improved medium for the isolation of bacteria from marine fish. Journal of General Microbiology, 52, 355–360. Sing, H., Can, A., Eustice, R., Lerner, S., & Mcphee, N. (2004). Seabed AUV offers new platform for high-resolution imaging. Eos, 85, 289. Sivasankar, P., Sugesh, S., Vijayanand, P., Sivakumar, K., Vijayalakshmi, S., Balasubramanian, T., et al. (2013). Efficient production of L-asparaginase by marine Streptomyces sp. isolated from Bay of Bengal, India. African Journal of Microbiology Research, 7, 4015–4021. Staley, J. T., & Konopka, A. (1985). Measurement of in situ activities of nonphotosynthetic microorganisms in aquatic and terrestrial habitats. Annual Review of Microbiology, 39, 321–346. Sudha, S. S. (2009). Marine environment: A potential source for L-asparaginase producing microorganisms. The IUP Journal of Biotechnology, 3, 57–62. Tanaka, S., Robinson, E. A., Appella, E., Miller, M., Ammon, H. L., Roberts, J., et al. (1988). Structures of amidohydrolases. Amino acid sequence of a glutaminase– asparaginase from Acinetobacter glutaminasificans and preliminary crystallographic data for an asparaginase from Erwinia chrysanthemi. The Journal of Biological Chemistry, 263, 8583–8591. Thenmozhi, C., Sankar, R., Karuppiah, V., & Sampathkumar, P. (2011). L-Asparaginase production by mangrove derived Bacillus cereus MAB5: Optimization by response surface methodology. Asian Pacific Journal of Tropical Medicine, 4, 486–491. Thirunavukkarasu, N., Suryanarayanan, T. S., Murali, T. S., Ravishankar, J. P., & Gummadi, S. N. (2011). L-Asparaginase from marine derived fungal endophytes of seaweeds. Mycosphere, 2, 147–155. Tonin, G. S., & Sodek, L. (1990). Asparaginase, allantoinase and glutamine synthetase activities in soybean cotyledons grown in vitro. Phytochemistry, 29, 2829–2831.

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Warrell, R. P., Jr., Arlin, Z. A., Gee, T. S., Chou, T. C., Roberts, J., & Young, C. W. (1982). Clinical evaluation of succinylated Acinetobacter glutaminase–asparaginase in adult leukemia. Cancer Treatment Reports, 66, 1479–1485. Whitman, W. B., Coleman, D. C., & Wiebe, W. J. (1998). Prokaryotes: The unseen majority. Proceedings of the National Academy of Sciences, 95, 6578–6583. Yao, M., Yasutake, Y., Morita, H., & Tanaka, I. (2005). Structure of the type I L-asparaginase from the hyperthermophilic archaeon Pyrococcus horikoshii at 2.16 angstroms resolution. Acta Crystallographica, D61, 294–301. Yong, W., Zheng, W., Zhnag, Y., & Zhu, J. (2003). L-Asparaginase-based regimen in the treatment of refractory midline nasal/nasal-type T/NK-cell lymphoma. International Journal of Hematology, 78, 163–167. Yooseph, S., Sutton, G., Rusch, D. B., Halpern, A. L., Williamson, S. J., Remington, K., et al. (2007). The sorcerer II global ocean sampling expedition: Expanding the universe of protein families. PLoS Biology, 5, e16. Zalewska-Szewczyk, B., Andrzejewski, W., & Bodalski, J. (2004). Development of antiasparaginase antibodies in childhood acute lymphoblastic leukemia. Pediatric Blood & Cancer, 43, 600–602. Zengler, K., Toledo, G., Rappe, M., Elkins, J., Mathur, E. J., Short, J. M., et al. (2002). Cultivating the uncultured. Proceedings of the National Academy of Sciences, 99, 15681–15686.

CHAPTER TWO

Marine Microbes as a Potential Source of Cellulolytic Enzymes N. Trivedi*,†,1, C.R.K. Reddy*,†,2, A.M. Lali{ *Division of Marine Biotechnology and Ecology, CSIR-Central Salt and Marine Chemicals Research Institute, Bhavnagar, India † Academy of Scientific and Innovative Research (AcSIR), New Delhi, India { DBT-ICT Centre for Energy Biosciences, Institute of Chemical Technology, Mumbai, India 2 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Classification of Cellulases 3. Marine Microbes as a Source of Cellulases 4. Marine Bacterial Cellulases 5. Marine Fungal Cellulases 6. Marine Yeast Cellulases 7. Conclusions and Future Prospectives Acknowledgment References

28 30 31 31 35 36 37 37 38

Abstract Marine environment hosts the wide range of habitats with remarkably high and diverse microbial populations. The ability of marine microorganisms to survive in extreme temperature, salinity, and pressure depends on the function of multivarious enzyme systems that in turn provide vast potential for biotechnological exploration studies. Therefore, the enzymes from marine microorganism represent novel bio catalytic potential with stable and reliable properties. Microbial cellulases constitute a major group of industrial enzymes that find applications in various industries. Majority of cellulases are of terrestrial origin, and very limited research has been carried out to explore marine microbes as a source of cellulases. This chapter presents an overview about the types of marine polysaccharases, classification and potential applications of cellulases, different sources of marine cellulases, and their future perspectives.

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Present address: DBT-ICT Centre for Energy Biosciences, Institute of Chemical Technology, Mumbai, India.

Advances in Food and Nutrition Research, Volume 79 ISSN 1043-4526 http://dx.doi.org/10.1016/bs.afnr.2016.07.002

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2016 Elsevier Inc. All rights reserved.

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1. INTRODUCTION Oceans cover about 70% of the earth’s surface and contain 97% of our planet’s water. Marine habitats represent the largest reservoir of biodiversity of the planet (Bull, Ward, & Goodfellow, 2000). The microbial diversity of marine origin has developed cellular machinery to thrive well even in extreme conditions. Marine microbes tolerating salt concentration of 1.7 M, temperature stability of 80–108°C, and high pressure of 60 MPa have been already reported in the literature (Marhuenda-Egea & Bonete, 2002; Singh et al., 2014). These properties have impelled worldwide research to target marine microbes as a potential source of novel enzymes. Various enzymes have been characterized from marine microbes isolated from seawater and different marine sediments. The main marine enzymes reported so far includes lipases, proteases, laccases, and polysaccharases. The polysaccharide-degrading enzymes from marine microbes have gained global attention due to novel industrial applications. Apart from the cellulases (Trivedi et al., 2011a) and amylases (Chakraborty, Khopade, Kokare, Mahadik, & Chopade, 2009; Li et al., 2007; Mohapatra, Banerjee, & Bapuji, 1998), marine microbes have also known to produce seaweeddegrading enzymes such as agarases (Gupta, Trivedi, Gupta, Reddy, & Jha, 2013; Suzuki, Sawai, Suzuki, & Kawai, 2003), carrageenases (Lemoine, Collen, & Helbert, 2009; Sarwar, Matoyoshi, & Oda, 1987), alginate lyases (Huang et al., 2013; Kim et al., 2013), fucoidanase (Silchenko et al., 2013), ulvan lyases, etc. (Collen, Sassi, Rogniaux, Marfaing, & Helbert, 2011). The marine polysaccharases have been summarized in Fig. 1. Among all, cellulases have been studied extensively across the globe due to wide applications in fuel, leather, textile, agriculture, food, medical, paper, and pulp industries (Menendez, Garcia-Fraile, & Rivas, 2015; Trivedi et al., 2011b). Potential industrial uses of cellulases have been summarized in Fig. 2. Increasing energy securities and global warming issues due to emissions of greenhouse gases led to finding other renewable energy sources. Among all, plant biomass has been given high priority to be used as a source of transportation fuels (mainly bioethanol) and chemicals. Cellulosic-richbiomass has been well recognized as a major source of energy feedstock or as a raw material for production of high value chemicals (Cherry & Fidantsef, 2003; Perlack et al., 2005). Cellulose, the most abundant and renewable organic polymer, is a linear homopolymer of D-glucose linked

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Fig. 1 Different type of marine polysaccharases.

Fig. 2 Potential industrial uses of cellulases.

with 1,4-beta acetal bond and constitutes primary structural cell wall material in both lower and higher plants (Saha, Roy, Sen, & Ray, 2006). Cellulose has been mainly used to produce ethanol through fermentation. Among other plant cell wall polysaccharide, cellulose is the most

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recalcitrant polymer to catalytic degradation. Cellulose could be hydrolyzed to its monomer, ie, glucose, either by chemical (acid), or enzymatic route. Currently, chemical hydrolysis is the process most widely followed to produce sugars from cellulose. Due to combination of high temperatures and strong acids, chemical hydrolysis leads to the degradation and accumulation of nonsugar by-products such as 5-hydroxymethylfurfural, formic acid, levulinic acid, and acetic acid (organic acids) which pose problems in downstream process (Mussatto & Roberto, 2004). As an alternate, enzymatic hydrolysis represents the greener approach for the saccharification of cellulosic-rich biomass. Cellulase is an inducible enzyme that catalyzes the hydrolysis of cellulose. It belongs to the class of hydrolases and is mainly produced by fungi, bacteria, protozoa, insects, and termites (Ozioko, Ikeyi, & Ugwu, 2013). Microbes produce cellulolytic enzymes as a single unit or in the form of cellulosomes (Bayer, Lamed, & Himmel, 2007). Cellulase has been commercialized during 1960s but still has some shortfalls such as low enzyme activity and stability in certain ecosystems associated with high salt, strong acid, strong alkali, and low temperature. In the global enzyme market, cellulases are the third most important industrial enzymes (15%) after amylase (25%) and protease (18%) (Sajith, Priji, Sreedevi, & Benjamin, 2016). Recently, the research on cellulases has significantly been increased due to its role in bioethanol production from cellulosic-rich biomass. By 2035, the global ethanol demand has been projected to increase by 3.5-fold (Limayem & Ricke, 2012). The high cost of enzyme production is a major limiting factor in commercialization of bioethanol production through enzymatic hydrolysis. Therefore, exploration for finding new and potential sources of cellulase is always desirable for economical enzyme production at industrial scale. Microorganisms with biochemical diversity and possible genetic modification are targeted as a source of potential enzymes. Microbial cellulases isolated from marine environment show higher degree of stability in different harsh conditions such as pH, temperature, and salinity.

2. CLASSIFICATION OF CELLULASES Cellulose hydrolysis via enzymatic route is accomplished synergistically by actions of group of enzymes known as cellulases. Based on their hydrolysis potential, cellulases have been classified into three types (Bhat & Bhat, 1997; Narra, Dixit, Divecha, Madamwar, & Shah, 2012), namely:

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(1) Endo-1-4-β-glucanase (EG, EC 3.2.1.4): Breaks down glycosidic bond in amorphous region (2) Exocellulase/cellobiohydrolase (CBH, EC 3.2.1.91): attaches to crystalline ends of cellulose chain producing cellobiose (3) β-Glucosidase (BGL, EC 3.2.1.21): Breaks down glucosidic bond of cellobiose and cellodextrins into glucose (Bhat & Bhat, 1997) Till date, a majority of cellulase producing microbes have been isolated from terrestrial sources. Some efforts have been made to explore the marine-based sources for cellulase production which are summarized below.

3. MARINE MICROBES AS A SOURCE OF CELLULASES The major cellulases producing bacterial genera include Bacillus, Clostridium, Cellulomonas, Bacteriodes, Acetivibrio, Rumminococcus, Geobacillus, Alteromonas, Vibrio and fungal genera include Aspergillus, Trichoderma, Chrysosporium, Penicillium (Assareh, Zahiri, Noghabi, Aminzadeh, & Khaniki, 2012; Dotsenko, Sinitsyna, Hinz, Wery, & Sinitsyn, 2012; Nandakumar, Thankur, Raghavarao, & Ghildyal, 1994; Roboson & Chambliss, 1989). The cellulolytic activity and stability vary with the source of origin. The cellulases used in industries require higher activity and stability in order to endure diverse harsh conditions. Globally, several research groups have been involved in isolation and characterization of cellulase producing microorganism with higher catalytic activity and stability. In comparison to terrestrial environment, microbes from marine habitat with hyper variable conditions, such as high pH, temperature, pressure, salinity, oxidative stress, metals, radiations, and chemicals could represent the novel source of extracellular enzymes with higher catalytic potential (Dalmaso, Ferreira, & Vermelho, 2015). So far, cellulases with alkali-halotolerant, organic solvent stable, ionic liquid stable, thermostable characteristics have been reported from marine microbes (Rastogi et al., 2010; Trivedi et al., 2011a, 2011b; Trivedi, Gupta, Reddy, & Jha, 2013; Yu & Li, 2015). Table 1 summarizes various cellulases recently studied from marine microbial sources.

4. MARINE BACTERIAL CELLULASES Increasing demand for industrial enzymes with desired characteristics turned attention toward exploration of marine microbes as a potential source. Microorganism surviving in marine environment develops well-evolved

Table 1 Cellulases Most Recently Reported from Marine Microbes pH and Microorganisms Isolated From Temperature

Characteristics

References

Gracilibacillus sp. SK1

Salt lake

8 and 60°C

Alkali stable

Yu and Li (2015)

Aureobasidium pullulans 98

Sea saltern of yellow sea

5.6 and 40°C

Potential CMCase

Yanjun, Liang, Zhenming, and Xianghong (2015)

Bacillus cereus JD0404

Muddy sediments of mangrove swamps

7 and 50°C

Able to degrade agroresidues

Chantarasiri (2015)

Cladosporium sphaerospermum

Deteriorated seaweed Ulva

4 and 25°C

SSF derived

Trivedi, Reddy, Radulovich, and Jha (2015)

Brachybacterium, Brevibacterium, Seaweed Eucheuma Halomonas, Kokuria, Micrococcus, cottonii Nocardiopsis, Pseudomonas, and Streptomyces (genera)

4.8 and 50°C

Lignocellulosic biomass degradation

Santhi, Bhagat, Saranya, Govindarajan, and Jebakumar (2014)

Isoptericola sp. JS-C42

Marine sediments

7.6 and 30°C

Plant biomass degradation

Santhi, Gupta, Saranya, and Jebakumar (2014)

Bacillus VITRKHB

–

7.8 and 25.8°C Good industrial efficacy

Singh et al. (2014)

Bacillus sp. H1666

Seawater samples

7 and 50°C

Seaweed degradation

Harshvardhan, Mishra, and Jha (2013)

Bacillus halodurans CAS 1

Marine sediments

9 and 60°C

Thermostable and haloalkaline

Annamalai, Rajeswari, Elayaraja, and Balasubramanian (2013)

Pseudoalteromonas sp.

Sargassum polycystum (Brown seaweed)

5 and 45°C

Ionic liquid stable

Trivedi et al. (2013)

Aspergillus ZJUBE-1

Soil of sea

4–5 and 65°C

SSF derived

Liu, Xue, He, & Yao (2012)

Bacillus flexus

Deteriorated seaweed Ulva

10 and 45°C

Alkali-halotolerant

Trivedi et al. (2011a)

Bacillus aquimaris

Deteriorated seaweed Ulva

11 and 45°C

Organic solvent stable

Trivedi et al. (2011b)

Aspergillus terreus

Marine water

7 and 37°C

Cellulase production Padmavathi, Agarwal, and through agro-waste residue Nandy (2012)

Aspergillus niger

East China Sea

6 and 28°C

Cellulase production using Xue, Chen, Lin, Guan, and E. crassipes, raw wheat bran, Yao (2012) raw corn cob, raw rice straw

Chaetomium sp.

Leaves of the mangroves

9.7 and 50°C

Cellulase production using agricultural and industrial waste

Mucor plumbeus

Ravindran, Naveenan, and Varatharajan (2010)

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cellular machinery suitable for thriving in extreme conditions of pH, temperature, pressure, and salinity (Singh et al., 2014). Recently, Annamalai et al. (2013) reported an extracellular thermostable, haloalkaline cellulase from Bacillus halodurans CAS. The enzyme showed thermal stability at 80°C and pH stability at 12 along with compatibility with different detergents and organic solvents. Similarly, alkali stable cellulase from halophilic Gracilibacillus sp. SK1 has been used in saccharification of corn stover and rice straw for bioethanol production. The ethanol yield was found to be 0.186 g/g reducing sugar with conversion efficiency of 58.2% (Yu & Li, 2015). Apart from the cellulases of terrestrial origin, microbial cellulases from marine environment have also shown successful conversion of cellulose-based plant biomass to fermentable sugar. Bacillus cereus JD0404 isolated from the muddy sediments of mangrove swamps showed bioconversion of cellulose-based biomass (Chantarasiri, 2015). In another recent study, cellulases from Enhydrobacter sp. ACCA2 were used to hydrolyze different plant biomasses such as bamboo, cumbu leaf, cumbu stem, sorghum leaf, and sorghum stem (Premalatha, Gopal, Jose, Anandham, & Kwon, 2015). Recently, extensive work has been carried out in the field of algal biofuel using cellulases of marine origin. Trivedi et al. (2011a, 2011b, 2013, 2015) focused on isolation of different marine microbes with cellulolytic potential. Among the different marine microbes isolated, B. flexus produced alkali halotolerant cellulase with a recovery of 25.03% and purity fold of 22.31. The optimum pH and temperature were found to be 10 and 45°C, respectively. Another strain Pseudoalteromonas sp. was found to stable in six different ionic liquids. The activity measured at 5% (v/v) was maximum with 1-ethyl-3-methylimidazolium bromide ([EMIM]Br) followed by 1-ethyl-3-methylimidazolium acetate ([EMIM] Ac), 1-butyl-3-methylimidazolium chloride ([BMIM]Cl), 1-ethyl-3methylimidazolium methanesulfonate ([C2MIM][CH3SO3]), 1-butyl-3methylimidazolium trifluoromethanesulfonate ([BMIM][OTF]), and 1-butyl-1-methylpyrrolidinium trifluromethanesulfonate ([BMPL][OTF]) with 115%, 104.7%, 102.2%, 98.33%, 93.84%, and 92.67%, respectively, and >80% activity at 15% (v/v) in all ionic liquids (ILs). Industrial applicability of the cellulase was also evaluated by using green seaweed Ulva lactuca as a carbon source in medium containing different ionic liquids. The specific activity of cellulase in IL containing reaction medium was found to be twofold higher in comparison to aqueous-based reaction medium. Similarly,

Potential Source of Cellulolytic Enzymes

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Harshvardhan et al. (2013) also reported cellulase from marine Bacillus sp. H1666 showing saccharification of marine macrophytic green alga U. lactuca in single step. Apart from seaweeds, microalgae have also been hydrolyzed using cellulase for biogas production. Mun˜oz, Hidalgo, Zapata, et al. (2014) used cellulases from nine different marine bacteria belonging to the genera Aeromonas, Pseudomonas, Chryseobacterium, and Raoultella for the hydrolysis of cell walls of Botryococcus braunii and Nannochloropsis gaditana. The cellulases have also been reported from microbes isolated from Antarctic region. Ferres, Amarelle, Noya, and Fabiano (2015) studied cellulases from different bacterial genera such as Pseudomonas, Pyschrobacter, Pseudoalteromonas, Loktanella, Flavobacterium, Arthrobacter, Polaromonas, Rhodococcus, Cryobacterium, Janthinobacterium, etc. Microbes surviving in extreme conditions of desiccation and freezing got adapted by metabolic functions and the synthesis of structurally adapted enzymes. Recently, Deep, Poddar, and Das (2016) have successfully cloned, overexpressed, and characterized halostable, solvent-tolerant novel β-endoglucanase from a marine bacterium Photobacterium panuliri strain LBS5T. The enzyme is monomeric and has molecular weight of 53 kDa with an optimum pH and temperature of 4 and 40°C, respectively. Further, the structural analysis of recombinant β-endoglucanase showed presence of 25% helix, 30% sheets, and 56% irregularities. The enzyme showed stability in different organic solvents with an increase in enzyme activity by 1.5-fold. The recombinant enzyme showed stability in 50% v/v concentrations of solvents which was higher than cellulases from B. vallismortis RG-07 (30% v/v), Thalossobacillus sp. LY18 (20% v/v) and B. aquimaris (20% v/v) (Gaur & Tiwari, 2015; Li, Wang, Li, & Yu, 2012; Trivedi et al., 2011b). Extremophilic microorganisms, mainly thermophiles and alkaliphiles, are potential lignocellulolytic enzyme producers (Trincone, 2011). Marine thermophilic microorganisms, such as Pyrococcus sp. (Matsui et al., 2000) and Thermotoga sp. (Duffaud, McCutchen, Leduc, Parker, & Kelly, 1997), have been reported for glycoside hydrolases.

5. MARINE FUNGAL CELLULASES Cellulases isolated from marine fungus have higher saccharification potential over bacterial cellulases. Recently, Trivedi et al. (2015) reported solid state fermentation (SSF)-derived cellulase from Cladosporium sphaerospermum using green seaweed Ulva fasciata as substrate. The optimum

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enzyme production was attained at 25°C with 60% moisture level, pH 4 and incubation period of 4 days. The optimum CMCase and FPase activity were found to be 9.56  0.53 and 8.83  0.37 U/g DW of seaweed. The enzyme was found to be active and stable in the presence of different ILs, namely, [EMIM]Ac, [BMIM]Cl, [BMIM][OTF], and [BMPL][OTF]. At 10% v/v concentration, enzyme retained 72.17–85.04% activity in all the ILs. After 24 h of preincubation in all ILs (10% v/v), enzyme activity was in the range of 73.77–93.67%. The hydrolysis of algal biomass with SSF-derived cellulase was optimized. The hydrolysis of U. fasciata feedstock with enzyme (10 U/g) for 24 h at 40°C and pH 4 gave maximum reducing sugar yield of 112  10 mg/g DW which on fermentation gave an ethanol yield of 0.47 g/g reducing sugar corresponding to 93.81% conversion efficiency. The ethanol yield estimated under optimized conditions was 4.4 kg/100 kg dry algal biomass. Xue et al. (2012) demonstrated an environment-friendly process to produce cellulase (17.80 U/g DW) by a marine Aspergillus niger under SSF using seawater, raw E. crassipes, raw rice straw, raw corn cob, and wheat bran. Several efforts have also been made to produce cellulases using cheap alternative carbon sources such as agricultural and industrial wastes. Padmavathi et al. (2012) explored marine fungal strain for cellulase production using 10 agro-waste residues. Among all, eucalyptus was found to be good carbon source for the cellulase production (733.3 IU/mL) at optimum pH of 7 and temperature of 37°C. Similarly, Ravindran et al. (2010) optimized the cellulase production from marine fungi using cotton seed, sugarcane bagasse, rice bran, and waste paper as a carbon source. The enzyme showed high activity and stability under neutral to alkaline pH and high temperature.

6. MARINE YEAST CELLULASES Marine yeast has gained considerable attention as a source of novel enzymes. Most of the yeast-derived enzymes, namely, amylase, alkaline protease, acid protease, phytase, lipase, inulinase have been isolated from terrestrial microorganisms. These enzymes have various potential applications in food, pharmaceutical, mariculture, and fermentation industries. Similarly, marine yeasts could also play a major role as a source of novel enzymes due to their ability to tolerate extreme harsh conditions. Kudanga and Mwenje (2005) reported endoglucanase and exoglucanase production from some sp. of Aureobasidium pullulans, black yeast. Similarly,

Potential Source of Cellulolytic Enzymes

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Zhang and Chi (2007) isolated and screened 19 strains of marine yeasts for cellulase production and optimized the conditions for higher cellulase production. Recently, Yanjun et al. (2015) isolated marine yeast from surface seawater of sea saltern at Yellow sea (China) for cellulase production. The strain 98 showed higher cellulase production (CMCase 4.51 U/mg and FPAase 4.75 U/mg protein) and was identified as A. pullulans. The molecular mass of the purified CMCase from A. pullulans 98 was 67.0 kDa. This strain has ability to survive in different marine environments and therefore play crucial role in biodegradation of polymers in marine environment (Nagahama, 2006). Apart from cellulases, A. pullulans has been shown to produce different enzymes such as protease (Chi, Ma, Wang, & Li, 2007), lipase (Wang, Chi, Wang, Liu, & Li, 2007), and amylase (Li et al., 2007).

7. CONCLUSIONS AND FUTURE PROSPECTIVES Marine microorganisms are being intensively studied worldwide, although systematic knowledge of the physiology, genetics, metabolism, and enzymology is limited. Therefore, it is necessary to explore the marine microorganism for diverse and unique applications with systematic knowledge. With the developments in biotechnology, industrial cellulases market is expected to grow in near future. Recently, focus has been shifted from terrestrial sources to marine-based sources mainly microorganisms for the production of novel cellulases due to their ability to survive in harsh conditions. Cellulases isolated from these organisms have shown their potential for bioconversion processes which have role in bioenergy-based industries. The study of marine microorganisms for cellulases has been significantly strengthened using the synthetic and system biology approach. The major future challenges for cellulase production includes (1) isolation and screening of potential cellulase producers with high stability and activity under extreme conditions; (2) design of improved biomass pretreatment strategies for better cellulase interaction; (3) manipulation of genetic engineering techniques for over production of enzyme with higher activity, stability, and process tolerance; and (4) enzyme production cost.

ACKNOWLEDGMENT Mr. Nitin Trivedi gratefully acknowledges the Department of Scientific and Industrial Research (DSIR), New Delhi for awarding Research Fellowship.

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Harshvardhan, K., Mishra, A., & Jha, B. (2013). Purification and characterization of cellulase from a marine Bacillus sp. H1666: A potential agent for single step saccharification of seaweed biomass. Journal of Molecular Catalysis B: Enzymatic, 93, 51–56. Huang, L., Zhou, J., Li, X., Peng, Q., Lu, H., & Du, Y. (2013). Characterization of new alginate lyase from newly isolated Flavobacterium Sp. S20. Journal of Industrial Microbiology and Biotechnology, 40, 113–122. Kim, E. J., Fathoni, A., Jeong, G.-T., Jeong, H. D., Nam, T. J., Kong, I. S., et al. (2013). Microbacterium oxydans, a novel alginate- and laminarin-degrading bacterium for the reutilization of brown-seaweed waste. Journal of Environmental Management, 130, 153–159. Kudanga, T., & Mwenje, E. (2005). Extracellular cellulase production by tropical isolates of Aureobasidium pullulans. Microbiology, 51, 773–776. Lemoine, M., Collen, P. N., & Helbert, W. (2009). Physical state of kappa-carrageenan modulates the mode of action of kappa-carrageenase from Pseudoalteromonas carrageenovora. The Biochemical Journal, 419, 545–553. Li, H. F., Chi, Z. M., Wang, X. H., Duan, X. H., Ma, L. Y., & Gao, L. M. (2007). Purification and characterization of extracellular amylase from the marine yeast Aureobasidium pullulans N13d and its raw potato starch digestion. Enzyme and Microbial Technology, 40, 1006–1012. Li, X., Wang, H. L., Li, T., & Yu, H. Y. (2012). Purification and characterization of an organic solvent-tolerant alkaline cellulase from a halophilic isolate of Thalassobacillus. Biotechnology Letters, 34, 1531–1536. Limayem, A., & Ricke, S. C. (2012). Lignocellulosic biomass for bioethanol production: Current perspectives, potential issues and future prospects. Progress in Energy and Combustion Science, 38, 449–467. Liu, J., Xue, D., He, K., & Yao, S. (2012). Cellulase production in Solid-State Fermentation by marine Aspergillus sp. ZJUBE-1 and its enzymological properties. Advanced Science Letters, 16, 381–386. Marhuenda-Egea, F. C., & Bonete, M. J. (2002). Extreme halophilic enzymes in organic solvents. Current Opinion in Biotechnology, 13, 385–389. Matsui, I., Sakai, Y., Matsui, E., Kikuchi, H., Kawarabayasi, Y., & Honda, K. (2000). Novel substrate specificity of a membrane-bound beta-glycosidase from the hyperthermophilic archaeon Pyrococcus horikoshii. FEBS Letters, 467, 195–200. Menendez, E., Garcia-Fraile, P., & Rivas, R. (2015). Biotechnological applications of bacterial cellulases. AIMS Bioengineering, 2, 163–182. Mohapatra, B. R., Banerjee, U. C., & Bapuji, M. (1998). Characterization of a fungal amylase from Mucor sp. associated with the marine sponge Spirastrella sp. Journal of Biotechnology, 60, 113–117. Mun˜oz, C., Hidalgo, C., Zapata, M., et al. (2014). Use of cellulolytic marine bacteria for enzymatic pretreatment in microalgal biogas production. Applied and Environmental Microbiology, 80, 4199–4206. Mussatto, S. I., & Roberto, I. C. (2004). Alternatives for detoxification of diluted-acid lignocellulosic hydrolyzates for use in fermentative processes: A review. Bioresource Technology, 93, 1–10. Nagahama, T. (2006). Yeast biodiversity in freshwater, marine and deep-sea environments. The yeast handbook biodiversity and ecophysiology of yeasts (pp. 241–262). Berlin: Springer. Nandakumar, M. P., Thankur, M. S., Raghavarao, K. S. M. S., & Ghildyal, N. P. (1994). Mechanism of solid particle degradation by Aspergillus niger in solid state fermentation. Process Biochemistry, 29, 545–551. Narra, M., Dixit, G., Divecha, J., Madamwar, D., & Shah, A. R. (2012). Production of cellulases by solid state fermentation with Aspergillus terreus and enzymatic hydrolysis of mild alkali-treated rice straw. Bioresource Technology, 121, 355–361.

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Trivedi, N., Reddy, C. R. K., Radulovich, R., & Jha, B. (2015). Solid state fermentation (SSF)-derived cellulase for saccharification of the green seaweed Ulva for bioethanol production. Algal Research, 9, 48–54. Wang, L., Chi, Z. M., Wang, X. H., Liu, Z. Q., & Li, J. (2007). Diversity of lipase-producing yeasts from marine environments and oil hydrolysis by their crude enzymes. Annals of Microbiology, 57, 495–501. Xue, D. S., Chen, H. Y., Lin, D. Q., Guan, Y. X., & Yao, S. J. (2012). Optimization of a natural medium for cellulase by a marine Aspergillus niger using response surface methodology. Applied Biochemistry and Biotechnology, 167, 1963–1972. Yanjun, R., Liang, Z., Zhenming, C., & Xianghong, W. (2015). A carboxymethyl cellulase from a marine yeast (Aureobasidium pullulans 98): Its purification, characterization, gene cloning and carboxymethyl cellulose digestion. Journal of Ocean University of China, 14, 913–921. Yu, H.-Y., & Li, X. (2015). Alkali-stable cellulase from a halophilic isolate, Gracilibacillus sp. SK1 and its application in lignocellulosic saccharification for ethanol production. Biomass & Bioenergy, 81, 19–25. Zhang, L., & Chi, Z. M. (2007). Screening and identification of a cellulase producing marine yeast and optimization of medium and cultivation conditions for cellulase production. Journal of Ocean University of China, 37, 101–108.

CHAPTER THREE

Extremozymes from Marine Actinobacteria J. Suriya*, S. Bharathiraja†, M. Krishnan*, P. Manivasagan{, S.-K. Kim§,1 *School of Environmental Sciences, Bharathidasan University, Tiruchirappalli, Tamil Nadu, India † CAS in Marine Biology, Annamalai University, Porto Novo, Tamil Nadu, India { Marine Bioprocess Research Center, Pukyong National University, Busan, Republic of Korea § Marine Bioprocess Research Center; Specialized Graduate School Science & Technology Convergence, Pukyong National University, Busan, Republic of Korea 1 Corresponding author: e-mail addresses: [email protected]; [email protected]

Contents 1. Introduction 2. Extremozymes 2.1 Thermophilic Extremozymes 2.2 Psychrophilic Extremozymes 2.3 pH Stability of Extremozymes 2.4 Halophilic and Organic Solvent Active Extremozymes 2.5 Barophilic Extremozymes 3. Extremozymes from Marine Actinobacteria 4. Application of Microbial Enzymes for Research 5. Structural Elucidation of Extremozymes 6. Large-Scale Production of Extremozymes 7. Industrial Applications of Extremozymes 8. Conclusion Acknowledgments References

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Abstract Marine microorganisms that have the possibility to survive in diverse conditions such as extreme temperature, pH, pressure, and salinity are known as extremophiles. They produce biocatalysts so named as extremozymes that are active and stable at extreme conditions. These enzymes have numerous industrial applications due to its distinct properties. Till now, only a fraction of microorganisms on Earth have been exploited for screening of extremozymes. Novel techniques used for the cultivation and production of extremophiles, as well as cloning and overexpression of their genes in various expression systems, will pave the way to use these enzymes for chemical, food, pharmaceutical, and other industrial applications.

Advances in Food and Nutrition Research, Volume 79 ISSN 1043-4526 http://dx.doi.org/10.1016/bs.afnr.2016.08.001

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1. INTRODUCTION In recent years enzymes have replaced several chemical processes in many industries due to its ecofriendly and less energy-consuming properties. Enzymes reduce the by-product formation due to its high specificity toward substrate which prevents the generation of expensive raw material wastage. Specificity of enzyme also minimizes side effects of enzyme-based therapeutic systems. It is very difficult to operate industrial processes at mild conditions as well as to hydrolyze complex substrates. Henceforth, there is an urgent need for discovering new enzymes able to hydrolyze complex substrates under extreme environmental conditions. Innumerable microorganisms have been screened to obtain novel enzymes. Among microorganisms, marine microorganisms constitute a promising source for many novel biocatalysts. This chapter focuses on unique properties of marine microbial enzymes, their applications, and future prospects. The marine biosphere is one of the richest and unexplored biospheres in the earth. Marine organisms are abundant and play an important role in the marine environment. Marine microorganisms have distinct features due to their unique physiological and metabolic characteristics; henceforth, they are easily adapted to the extreme environments such as high or low temperature, acidic or alkali environment, and low nutritional content found in the deep sea. These unique features have attracted many scientists to explore in depth. Marine microorganisms were already proven as a potential agent for the production of many metabolites including industrial enzymes (Chatellier, Bhagat, Karthik, Jaffar Hussain, & Jayaprakashvel, 2011). For the enzyme-based industrial reaction conditions, the enzymes should be stable and active at those conditions. Many marine microbial enzymes revealed such appropriate properties, proving its suitability for the different industrial processes, biotechnological research organic synthesis, and biomedical applications (Hassanein, Kotb, Awny, & El-Zawahry, 2011; Li et al., 2007; Zhu, Malik, & Hua, 2006). The potential of marine microbial enzymes to withstand extreme pH, temperature, organic solvents, and high salt concentration makes them available for many industrial processes (Lundberg et al., 1991; Plisova et al., 2005; Zhu et al., 2006). Several marine extremozymes have been exploited for biotechnological research, whereas others are exploited in pharmaceutical industry, biofuel industry, and textile industry (Lundberg et al., 1991; Verenium-Fuelzym, 2012). Pharmaceutically essential stereoselective compounds are obtained from solvent-tolerant

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marine microbial enzymes. Cold-adapted xylanases secreted by Antarctic microorganisms have been utilized in baking industry (Collins et al., 2006) and cold-active β-galactosidase produced by a marine psychrophilic bacterium that hydrolyzed 80% of the lactose in raw milk (Ghosh, Pulicherla, Rekha, Raja, & Sambasiva Rao, 2012; Pulicherla et al., 2013). Van de Voorde, Goiris, Syryn, Vanden Bussche, and Aerts (2014) obtained a lactose-hydrolyzing psychrotolerant β-galactosidase from the Antarctic marine bacterium Pseudoalteromonas haloplanktis and found potential application in tagatose production.

2. EXTREMOZYMES Extremophiles are organisms which have the potential to survive in the extreme conditions of life. At the time of evolution, marine microorganisms underwent extreme environmental challenges and adapted their metabolic pathways to survive in extreme environments resulting in the production of unique metabolites, which are not synthesized by terrestrial microorganisms. Extremophiles are usually the primary source of extremozymes. Marine enzymes have the potential to withstand extreme temperature or pH. Some are stable and active at extreme saline condition and can work in the presence of different chemicals and organic solvents. Stereochemical properties of marine enzymes also increase its value in commercial industrial processes. In some cases, a single enzyme has the combination of two or more above said features which is pivotal for the development of unique bioprocesses. For example, thermostable alkaliphilic proteases have been used in laundry detergents and these enzymes have the potential application in dry-cleaning industries if they are active and stable in the presence of organic solvents. Likewise thermostable solvent-tolerant enzymes have utilized in the biofuel industries and these enzymes can be exploited for biofuel production from seaweed biomass if they can work in saline environment.

2.1 Thermophilic Extremozymes Extremozymes have attracted more attention than other enzymes due to its potential to carry out the reaction at extreme temperatures which are usually not possible at ambient temperature. Most of the chemical reactions are carried out at high temperature because it increases the solubility of substrates, reaction rate, better mixing, and also reduces the viscosity and microbial contamination. In addition to this, it is very easy to purify the thermostable enzymes by heat treatment. Thermostable amylases proteases, lipases,

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cellulases, chitinases, carragenases, xylanases, polymerases, and oxidases have innumerable applications in food processing, detergent, paper, textile, biofuel industries, and biotechnological research. Bacillus, Thermus, Thermotoga, Thermococcus, Pyrococcus, and Sulfolobus families are mostly screened for the production of thermostable marine microbial enzymes. Moreover, thermostable archaeal enzymes are more stable at high pressure, and can exert proteolytic degradation in the presence of detergents and organic solvents (Egorova & Antranikian, 2005). Deep-sea hydrothermal vents are the major source for thermostable enzymes of hyperthermophilic archaea (Cornec, Robineau, Rolland, Dietrich, & Barbier, 1998; Hung et al., 2011; Legin, Ladrat, Godfroy, Barbier, & Duchiron, 1997). Pyrococcus abyssi produces a crude esterase keeping the initial activity at 90°C after 8:5 h incubation and also has a long half-life at 99°C after 22 h of incubation (Cornec et al., 1998). A thermostable amylase from Rhodothermus marinus produces branched glucan from amylase and shows maximum activity at 80°C (Seong-Ae, Ryu, Lee, & Moon, 2008). Thermus aquaticus and Pyrococcus furiosus are the two thermophilic archaea which produce Taq polymerase and Pfu polymerase, respectively. These are two mostly used thermostable DNA (deoxyribonucleic acid) polymerases in biotechnological research isolated from two thermophilic archaea (Chien, Edgar, & Trela, 1976; Lundberg et al., 1991). The P. furiosus was isolated from a geothermally heated marine sediments with temperatures in between 90°C and 100°C. Thermophilic esterase from Archaeoglobus fulgidus is cloned and overexpressed in Escherichia coli (Chakraborty, Khopade, Kokare, Mahadik, & Chopade, 2009). It shows the maximum activity at 70°C and pH 7.1. Thermococcus hydrothermalis produces thermostable α-amylase enzyme. The gene encoding for this enzyme has been cloned and expressed in E. coli (Hayashi, 1996). It is optimally active at 75–85°C and at pH 5.0–5.5. Thermococcus aggregans synthesized hyperthermostable glycosidase enzyme with pullulanase activity at 90°C that was cloned and expressed in E. coli (Michels & Clark, 1997). This enzyme is able to hydrolyze α-1,6- and α-1,4-glycosidic linkages in pullulan, resulting in the production of maltose, glucose, panose, and maltotriose. The enzyme has the ability to hydrolyze starch, amylose, and amylopectin, producing maltotriose and maltose.

2.2 Psychrophilic Extremozymes Cold-adaptive enzymes are most preferred in industries because it avoids degradation or evaporation of reaction components and it also decreases

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corrosion of metallic reactors. Mild heat treatment can easily inactivate coldactive enzymes; this property is useful in multistep process if the enzyme has undesired effect in a subsequent reaction step. In the textile industry cellulase enzymes are used for stonewashing. However long-term enzyme activity also leads to the loss of cotton fiber’s mechanical resistance. Many psychrophilic marine microorganisms are screened for obtaining cold adaptive including the members of the genus Aquifex, Alteromonus, Bacillus, Psychrobacter, and Methanococcus. Arctic and Antarctic microorganisms are major sources of cold-active enzymes. A cold-adaptive alkaline phosphatase enzyme that was isolated from an Antarctic bacterium revealed application in molecular biology research (Kobori, Sullivan, & Shizuya, 1984) and it is easily inactivated by mild heat treatment (55°C) that eliminates the need of separating the DNA after alkaline phosphatase treatment. Many cold-active enzymes have potential applications in baking industry, bioremediation, and as cold-wash additive (Collins et al., 2006; Jeon et al., 2009; Trincone, 2011). Amylase-producing gene of marine bacterium Zunongwangia profunda was cloned and overexpressed in E. coli retaining 39% of activity at 0°C and with potential application in detergent industry (Qin, Huang, & Liu, 2014). A cold-adaptive β-galactosidase produced by a marine psychrophilic bacterium hydrolyzed 80% of lactose in raw milk at pH 6.5 and temperature of 20°C (Ghosh et al., 2012; Pulicherla et al., 2013). Van de Voorde et al. (2014) obtained a lactose-hydrolyzing psychrotolerant β-galactosidase from the Antarctic marine bacterium P. haloplanktis and found potential application in tagatose production. Textile and paper industries utilize cold-adaptive polysaccharolytic enzymes. Cold-active lipases were isolated from Antarctic deep-sea sediments metagenome and also from bacterial species such as Pseudomonas and Psychrobacter (Zhang, Lin, & Zeng, 2007; Zhang & Zeng, 2006, 2008). These enzymes are highly active and stable at very low temperature; however, it lost its activity at ambient temperature. A cold-active lipolytic enzyme was obtained from Southern Ocean deep sea that retains 37% of its maximum activity at 0°C (Lin, Yang, Bian, & Huang, 2003). Cold-adaptive lipases are used for the bioremediation of fat-contaminated aqueous systems (Trincone, 2011). Pseudoalteromonas arctica produces cold-active esterase which is optimally active at 25°C and retains 50% of its maximum activity at 0°C (Khudary, Venkatachalam, Katzer, Elleuche, & Antranikian, 2010). It digests large numbers of nonsteroidal antiinflammatory drugs including ibuprofen. A cold-active xylanase was isolated from marine fungi associated with marine sponge (Del-Cid et al., 2014).

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2.3 pH Stability of Extremozymes Most of the enzymes are active at neutral pH and many enzymes are inactivated at extreme acid or basic environments. Several industrial reactions require acidic or alkaline conditions where most enzymes are unstable and inactivated—such reactions are only catalyzed by enzymes that can withstand extremely low or high pH, respectively. Many marine microorganisms have the ability to thrive at extreme acidic or alkaline conditions and enzymes derived from these microorganisms are also active at extremely low and high pH, respectively. Alkaliphilic protease, lipase, amylase, and cellulase are consumed by detergent industry as additive and also these enzymes are used for cleaning of contact lens. Thermotolerant alkaliphilic proteases of marine microbial origin are exploited as cleansing enzyme (Gouda, 2006; Greene, Griffin, & Cotta, 1996; Kumar, Joo, Koo, Paik, & Chang, 2004). Among alkaliphilic enzymes, alkaline phosphatases are widely used for industrial purposes (Plisova et al., 2005; Sebastian & Ammerman, 2009). Alkaline phosphatase was isolated from marine microorganisms which has potential application in biotechnological research and it was inactivated by mild heat treatment (at 55°C; Kobori et al., 1984). Food and pharmaceutical industries utilize acidophilic digestive enzymes. Marine Bacillus is the major source of several alkaliphilic enzymes while acidophilic enzymes are produced by marine fungi. Thermostable alkaline proteases of marine Bacillus are active in the presence of surfactants and bleaching agents; henceforth, they are used as additive in commercial detergent (Abou-Elela, Ibrahim, Hassan, Abd-Elnaby, & El-Toukhy, 2011; Gouda, 2006; Kumar et al., 2004). An alkaline protease reported from marine shipworm bacterium efficiently degrades lysozyme (Kumar et al., 2004) and it is resistant to hydrogen peroxide which is consumed to sterilize contact lenses. Two thermotolerant alkaliphilic amylases from marine microorganisms are resistant to surfactants and detergents and exploited by detergent industry (Chakraborty et al., 2011, 2009).

2.4 Halophilic and Organic Solvent Active Extremozymes High salinity is the characteristic feature of the marine environment and enzymes produced by marine microorganisms are reasonably stable and active in the presence of high salt concentration. Usually, salts are present in reaction systems as contaminant, reactant, or product; sometimes, salts are added to the bioprocess to stabilize proteins. Salt-active enzymes can be exploited to catalyze high salt content reactions where most common

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industrial enzymes cannot work. Halobacterium salinarum produces a chitinolytic extremozyme that is stable and active up to 1.5 M NaCl concentration and well suited for the decomposition of marine chitincontaining wastes (Sana, Ghosh, Saha, & Mukherjee, 2007). For fish/meat processing and peptide synthesis halotolerant proteolytic enzymes are used. Salt-tolerant polysaccharolytic enzymes are employed for the biofuel production from marine microalgae and seaweed biomasses. Saccharification of marine microalgae is carried out by a halotolerant amylase enzyme of marine bacterium in the presence of high NaCl concentration (Matsumoto, Yokouchi, Suzuki, Ohata, & Matsunaga, 2003). These enzymes have the potential to work at low water availability and some of them could not work in the absence of high salt concentration. Marine Streptomyces sp. D1 produces an amylase which was 100% active with 7% (w/v) NaCl after 48 h incubation, but its activity was suddenly decreased after 48 h incubation without NaCl (Chakraborty et al., 2009). Several halotolerant enzymes are also active in the presence of organic solvents (Marhuenda-Egea & Bonete, 2002; Sellek & Chaudhuri, 1999). Marine Aeromonus hydrophila produces an organic solvent-tolerant lipase and its production is highly dependent on NaCl concentration. Stability of these enzymes toward high salt and organic solvents makes this enzyme to be consumed for certain applications. Organic solvent-active enzymes catalyze reactions in biphasic mixtures and they are employed for chirally pure drug molecules synthesis. P. furiosus produces an organic solvent-active alcohol dehydrogenase which reduces ketones with low solubility in aqueous buffers (Zhu et al., 2006). This enzyme is highly resistant to iso-propanol, DMSO, methyl tert-butyl ether, and hexane and its activity was increased at elevated temperature without changing its enantioselective potential. Some of the organic solvent-active esterases/lipases have been obtained from marine microorganisms and utilized for biodiesel production and chirally pure compounds synthesis (Neelambari, Vasanthabharathi, Balasubramanian, & Jayalakshmi, 2011; Sana et al., 2007).

2.5 Barophilic Extremozymes Sign and magnitude of the activation volume determine the impact of pressure on bioprocesses. To carry out the reactions at high pressure, it is pivotal to use barophilic enzymes. These enzymes are mostly derived from Deepsea piezophilic microorganisms. Pressure-stable enzymes are mostly consumed in food processing industries for processing and the sterilization of food

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materials (Hayashi, 1996). Some of the barophilic enzymes from marine microorganisms have been studied for their activity, structure, flexibility, and potential applications in biotechnological research (Abe & Horikoshi, 2001). Deep-sea fungus reported from 5000 m depth of Central Indian Basin produces a barophilic enzyme which is stable and active at pressures as high as 300 bar (Damare, Raghukumar, Muraleedhran, & Raghukumar, 2006). Deep-sea bacterium Methanococcus jannaschii synthesizes a thermostable barophilic enzyme that is the best example for extremozymes of marine microorganisms (Michels & Clark, 1997). Because pressure regulates the enzyme activity and thermostability in this species. So, enzyme activity and thermostability was elevated by 3.4- and 2.7-fold, respectively, by 500 atm at 125°C. Similarly, hydrogenase also revealed the same behavior and its activity was enhanced by threefold when pressure was increased from 7.5 to 260 bars at 86°C (Shah & Clark, 1990).

3. EXTREMOZYMES FROM MARINE ACTINOBACTERIA Nowadays, researchers have discovered several novel enzymes from marine organisms such as bacteria, fungi, and actinomycetes that showed medicinal as well as industrial applications. Among these microorganisms, actinomycetes reveal immense potential in the production of extremozymes. Marine actinomycete Saccharopolyspora sp. A9 produces an amylase that is optimally active at pH 11.0 and also stable at pH 8.0–12.0; temperature of 55°C (Chakraborty et al., 2011). Chakraborty et al. (2009) also found that the amylase production in marine actinomycete Streptomyces sp. D1 was not depending on the growth of the actinomycete. A cold-active α-amylase was obtained from Nocardiopsis aegyptia isolated from marine sediment in Egypt (Abou-Elela, El-Sersy, & Wefky, 2009). It was optimally active at 25°C under acidic conditions. The isolation and immobilization of a thermotolerant α-amylase of haloalkaliphilic marine isolate Nocardiopsis sp. B2 was carried out by Chakraborty et al. (2014). Chakraborty, Raut, Khopade, Mahadik, and Kokare (2012) accessed the calcium-independent, thermostable α-amylase from haloalkaliphilic marine Streptomyces A3. This enzyme was stable and active in the presence of 10% NaCl, pH 9.0, and retained 40% of its activity at 90°C. Ramesh, Rajesh, and Mathivanan (2009) isolated 208 marine actinomycetes from the Bay of Bengal, India. Streptomyces spp. occupied 88% of total actinobacteria. Isolates 68, 72, 116, 157, 183 only produced amylase, cellulose, gelatinase, caseinase, and lipase, respectively. An alkaline amylase that revealed activity at pH 9.5,

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50°C, and 9% NaCl was isolated from Streptomyces rochei BTSS 1001 (Acharyabhatta, Kandula, & Terli, 2013). Thumar and Singh (2007) obtained an alkaline protease from halotolerant and alkaliphilic Streptomyces clavuligerus strain. This enzyme showed optimal activity at pH 9 and 5% NaCl. An alkaliphilic and salt-tolerant actinomycete, S. clavuligerus strain, was isolated from Western coast of India. It revealed optimal alkaline protease activity (130 U/mL) at pH 9.0 and 5% NaCl during the early stationary phase. Sucrose and gelatine were found to be good carbon and nitrogen sources for maximum protease production (Thumar & Singh, 2007). Organic solvent tolerance of this enzyme was also studied by Thumar and Singh (2009) and found that the butanol increased the enzyme production by 50-fold and it showed optimal activity at 70°C with acetone and ethanol. The enzyme was stable and active in the presence of xylene, acetone, and butanol. However, ethanol and benzene affected the catalysis of this protease. Three hundred marine samples were collected from different locations of Bay of Bengal for the isolation of 208 marine actinomycetes isolates. Among those strains, Streptomyces sp. MA1-1 revealed optimal alkaline protease activity at pH 9.0 and 50°C (Guravaiah, Prabhakar, Prameela, Santhoshi Kumari, & Guravaiah, 2010). Fulzele, DeSa, Yadav, Shouche, and Bhadekar (2011) screened nine different isolates from marine samples of the Indian Ocean for alkaline protease production. Marinobacter sp. produced maximum protease production compared to others. The crude enzyme of Marinobacter sp. was stable in the pH range of 5.0–9.0 and temperature 30–80°C. Hames and Uzel (2007) optimized the production of alkaline protease of marine alkaliphilic actinomycete MA1-1 using various carbon and nitrogen sources. An alkaline protease was purified from S. clavuligerus (Moreira, Albuquerque, Teixeira, Porto, & Filho, 2002). Mehta, Thumar, and Singh (2006) obtained an extracellular serine protease produced by alkaliphilic actinomycetes. An alkaline protease gene of two salt-tolerant actinomycetes was cloned and expressed in E. coli strain by Gohel and Singh (2012a). Gohel and Singh (2012b) purified and characterized a thermostable alkaline protease from a salt-tolerant alkaliphilic actinomycete, Nocardiopsis alba OK-5. A thermostable alkaline serine protease was produced by halotolerant alkaliphilic actinomycetes Brachystreptospora xinjiangensis OM-6 that was stable at 80°C and also resistant to surfactants, urea denaturation, reducing and oxidizing agents (Gohel & Singh, 2013). The thermodynamics of a Ca2+-dependent thermostable alkaline protease of haloalkaliphilic actinomycetes N. alba OK-5 was studied (Gohel & Singh, 2015). The 20-kDa enzyme revealed optimal activity at 70°C in

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0–3 M NaCl and also stable in the presence surfactants, cations, H2O2, β-mercaptoethanol, and commercial detergents. Abdel-Aziz, Hamed, Ghaly, and El-Shafei (2012) showed the optimal activity of alkaline protease produced from marine Streptomyces albidoflavus in the pH range of 8–11. Nocardiopsis dassonvillei NCIM 5124 was isolated from oil-polluted marine sediment and produced two types of alkaline serine endopeptidases (Dixit & Pant, 2000). Ramesh et al. (2009) isolated 191 marine actinomycetes from 256 marine samples. Among those strains, Streptomyces fungicidicus produced the high thermostable alkaline protease enzyme. Purification and characterization of 40 kDa alkaline serine protease of Haloalkaliphilic bacterium sp. AH-6 was carried out by Dodia et al. (2008). Suthindhiran, Jayasri, Dipali, and Prasar (2014) reported a 22-kDa thermostable alkaline protease from marine Actinopolyspora sp. VITSDK2. Dasilva, Yim, Asquieri, and Park (1993) reported the alkaline cellulase from alkaliphilic Streptomyces sp. S36-2 that was optimally active at pH 6.0–7.0 and 55°C even though an alkaliphilic Streptomyces KSM-9 produced an alkaline cellulase (Damude et al., 1993). Dasilva et al. (1993) also found the optimal activity at pH 8–9. An endocellulase-produced gene of alkaliphilic Streptomycete was cloned and expressed in E. coli strain to get maximum production (van Solingen et al., 2001). Gene responsible for the production of extracellular alkaline lipase of S. rimosus R6-554W was cloned and overexpressed in a lipase-deficient S. rimosus that produced 22-fold higher activity than the original strain (Vujaklija, Schroder, & Abramic, 2002). Thermostable lipase produced by marine Streptomyces sp. W007 was expressed in Pichia pastoris X-33, and it was biochemically characterized (Yuan, Lan, Xin, Yang, & Wang, 2016). Nocardiopsis albus subsp. prasina OPC-131 an alkaliphilic actinomycete produced two types of chitinases with optimal activity at pH 8 and 7, respectively (Tsujibo, Yoshida, Miyamoto, Hasegawa, & Inamori, 1992). A thermostable chitinase of Streptomyces thermoviolaceus OPC-520 was overexpressed, purified, and characterized by Christodoulou, Duffner, and Vorgias (2001). An exoinulinase was obtained from Nocardiopsis sp. DN-K15 isolated from marine sediments in China (Lu, Li, & Guo, 2014). The enzyme was thermostable and alkali tolerant and optimally active in a wide range of pH (5.0–11.0). For L-asparaginase production, Saleem Basha, Rekha, Komala, and Ruby (2009) isolated 10 marine actinomycetes from marine sediment samples. S3 isolate showed maximum activity (65 μg/mL) than others at 50°C and pH 7.5. Polyhydroxybutyrate depolymerase has been reported from N. aegyptia from marine seashore sediments in Egypt (Ghanem, Mabrouk, Sabry, & El-Badan, 2005).

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Actinomycetes produced an unusual thiol compound for its protection against variety of challenges known as mycothiol (MSH). Henceforth, the enzymes involved in the MSH-dependent metabolism are targets for new drug discovery against Mycobacterium tuberculosis and other infectious actinomycetes (Vignesh, Raja, & James, 2011).

4. APPLICATION OF MICROBIAL ENZYMES FOR RESEARCH Most extracellular enzymes of marine microorganisms are exposed to extreme conditions; henceforth, several extremozymes are extracellular enzymes. Previously we could only find research articles with physicochemical characterization of crude, partially purified, or purified enzymes. In recent years, cloning and overexpression of marine-based enzymes in particular extremozymes is a common practice due to the development of genetic engineering tools. Now, many marine microbial enzymes have been engineered to obtain the desired characteristics and structures of a few marine enzymes (Choi et al., 2011; Hatada et al., 2006; Sakaguchi et al., 2004). Crucial proteins are screened from uncultivable marine microorganisms by using metagenomic analysis. In addition to this, media composition and fermentation parameters are optimized using mathematical models. Enzymes of marine microbial origin have other nontraditional applications including waste management and biofuel production (Basheer et al., 2011; Khandeparker, Verma, & Deobagker, 2011; Menon, Mody, Keshri, & Jha, 2010). In the past, only a particular group of scientists worked on extremozymes. Today, it has been studied widely due to the development of molecular biology and recombinant DNA technologies. It is not economic to produce an enzyme in large scale from a native host owing to its low productivity and nutrient wastage. Usually, expensive media components and distinct fermentation conditions have to be provided for marine microorganisms to get the enzyme of interest. These drawbacks are overcome by cloning and expression of gene of interest from marine microorganisms in the expression host. It is also easy and efficient to obtain purified product from expression host than from the complex natural host. Among marine microbial enzymes, thermotolerant DNA polymerases are widely utilized in research. Thermococcus litoralis and P. furiosus produce Vent polymerase and Pfu polymerase, respectively. These two marine microbial enzymes have been mostly utilized for PCR due to their stability

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at >95°C. Tth DNA polymerase was obtained from Thermus thermophile which has reverse transcriptase activity in the presence of Mn2+ ion and also revealed optimal activity at 70–80°C. Henceforth, it is applied for one-step RT-PCR and real-time PCR. Biotec Pharmacon has marketed salt-active nucleases, thermolabile DNAses, and uracil DNA glycosylase for research. Thermostable α-amylase of deep-sea microorganism was discovered by Vernium Corporation (Verenium-Fuelzym, 2012). The enzyme with brand name Fuelzyme is highlighted as a next generation for starch liquefaction. α-Amylase, which is active in a wide range of temperatures, hydrolyzes α-1,4 linkage of starch, and yields dextrin and oligosaccharides, and it is utilized for fuel ethanol production. Marine fungus Candida antarctica produces two lipases namely C. antarctica Lipase A and Lipase B applied for the transesterification production of biodiesel (Mahmoudian, Eddy, & Dowson, 1999). Yarrowia lipolytica is used as the expression system for cloning and expression of alkaline protease gene of Aureobasidium pullulans 10 and A. pullulans HN2-3. A. pullulans 10 gene was expressed as an extracellular protein and easily purified from the culture medium, whereas gene of A. pullulans HN2-3 was expressed as surface-displaying alkaline protease (Ni et al., 2009), which produces bioactive peptide that has angiotensin-converting enzyme inhibitory activity and antioxidant activity. Several bioactive peptides are produced from marine enzymes using different raw materials. Two alkaline serine protease-encoding gene of marine Alteromonas sp. strain O-7 was cloned and overexpressed in E. coli (Tsujibo et al., 1996, 1993). Alkaline protease gene of marine fungus Engyodontium album was also cloned in E. coli (Jasmin et al., 2010). PCR-based screening was used to know the sequence of a Psychrobacter sp. lipase gene by primers retrieved from multiple sequence alignments of prokaryotic lipases group (Parra et al., 2008). This gene was cloned into expression vector and overexpressed in E. coli as a fusion protein retained high lipolytic activity and also revealed psychrophilic property with optimum activity at 20°C. Site-directed mutagenesis was used to study the oxidative stability of an α-amylase (Hatada et al., 2006). This enzyme was produced by deep-sea marine Bacillus and is stable and active at 1 M H2O2 after 1 h incubation. Mutagenesis study revealed that the replacement of methionine residue by nonoxidizable Leu198 is responsible for the hyperoxidative stability of this enzyme. Metagenomic approach has been used for discovering many marine enzymes especially extremozymes. Metagenomic library was constructed from Baltic Sea sediment bacteria using 47,000 clones by extracting the

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prokaryotic DNA and cloned in pCC1FOS plasmid. The size of the insert ranges from 24 to 39 kb. A marine lipase was screened from this library by functional screening. Screened clones were subcloned and sequenced. Sequence results revealed that it was a 978-bp lipase and its molecular mass was 35.4 kDa that showed 54% amino acid similarity with a Pseudomonas putida esterase (Hardeman & Sjoling, 2007). Two novel esterase genes were screened by activity-based screening from marine microbial metagenomic library (Chu, He, Guo, & Sun, 2008). These esterases had 277 and 328 amino acids which were purified and biochemically characterized. The esterase which has 328 amino acids was highly stable in the presence of organic solvents. Another novel esterase from marine sediment microbial metagenomic library was cloned, overexpressed, and characterized by Xu, Hu, Yuan, and Zhu (2010). The screened protein was a psychrophilic esterase which showed 71% sequence similarity to another marine sediment metagenome esterase. A metagenomic library was constructed for marine sponge Hyrtios erecta associated bacteria to screen esterase enzyme (Okamura et al., 2010), and E. coli cells were used for cloning and expression of this enzyme getting it thermostable and halotolerant. Marine metagenomic library was used for screening α-amylase-encoding gene. It was cloned in pUC19 vector and overexpressed in E. coli host cells (Liu et al., 2012). It revealed 50–72% sequence similarity to four putative glycosidases of four different marine bacteria.

5. STRUCTURAL ELUCIDATION OF EXTREMOZYMES Innumerable works have been done on the characterization of marine microbial enzymes but only few reports were available on structural studies of these enzymes. It is essential to elucidate the 3D structure of enzymes to know the relationship between structure and activity and it is also helpful to improve the protein activity by protein engineering. Substrate specificity of enzymes is determined from catalytic domain structures and other domains are useful for determining other features required for enzyme activity such as ligand binding and stability of the enzyme. Catalytic domain of a β-1,3-xylanase of marine bacterium Vibrio sp. AX-4 was studied with the help of crystallization and preliminary X-ray analysis (Sakaguchi et al., 2004). Another enzyme used for the structural studies is methanol dehydrogenase of the marine bacterium Methylophaga aminisulfidivorans MPT (Choi et al., 2011).

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Chinese Yellow Sea Flavobacterium YS-80 sp. and Antarctic Pseudomonas sp. produce psychrophilic alkaline proteases showing a homologous structure (Aghajari et al., 2003; Zhang et al., 2011). N-terminal domain is responsible for proteolytic activity of the enzyme whereas C-terminal domain determines the stability of protein structure. Zn2+ and Ca2+ ions are combined with crystal structure of these proteins. Alkaline protease of Yellow Sea Flavobacterium is highly thermotolerant and has a rigid structure compared to Antarctic Pseudomonas protease. These properties are due to the respective habitat of the enzyme-producing microorganisms and it is also used for shedding light on the structure–environment adaptation of these enzymes.

6. LARGE-SCALE PRODUCTION OF EXTREMOZYMES Major challenges for commercialization of several marine enzymes are the continuous supply of sufficient quantity of the product and cost of largescale production. Optimization of media components and fermentation parameters such as pH, temperature, aeration, and pressure is crucial for large-scale production of marine microbial enzyme. It is very difficult to cultivate marine microorganisms because it requires unique media composition and physical conditions of their native habitat. For economical production of marine enzymes inexpensive raw materials are used in solid-state and submerged fermentations. Addition of NaCl and trace metals or sea water increases marine enzyme productivity. A 2-L fermenter was used for optimization of fermentation parameters and media components for protease production by the marine yeast, A. pullulans (Chi, Ma, Wang, & Li, 2007) and the production was maximum with initial pH 6.0 after 30 h incubation at 24.5°C. Cost-effective and easily available raw materials were optimized for large-scale production of thermostable alkaline protease from marine Bacillus sp. (Kumar et al., 2004), obtaining maximum activity in a soybean–casein medium, with pH 9.6 at 42°C after 40 h of fermentation of alkaline protease production from marine bacterium Teredinobacter turnirae (Elibol & Moriera, 2005). Had the highest maximum production with 2.5% inoculums level, 1% soybean concentration at pH 7.34. Agitation and aeration rates were optimized for extracellular protease production from marine Vibrio harveyi sp. (Estrada-Badillo & MarquezRocha, 2003). At 700 rpm and 0.5 vvm air flow they got the maximum protease production and also observed that the addition of skim milk powder in Zobell medium elevated the enzyme activity by fivefold. Agitation

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determines the growth rate of the culture and mostly stationary phase induces the expression of several enzymes. Response surface methodology was used for esterase production by marine Bacillus licheniformis MP-2 strain (Ping et al., 2008). Eleven parameters essential for esterase production were studied with Plackett–Burman design. Production of enzyme was severely influenced by the amount of soybean cake, peanut cake, and inoculum volume. Same approach was used for optimization of media components and fermentation parameters for a cold-adapted amylase production from marine bacterium Wangia sp. 52 (Liu et al., 2011). At optimized conditions, they obtained 10-fold increase of the amylase production than control shake flask culture. Gene responsible for thermostable esterase production of thermophilic marine Pyrococcus sp. was cloned and overexpressed in E. coli (Ravot et al., 2004). Then overexpressed protein was purified and used for industrial applications as a result of high product recovery yield (>70%) and enzyme concentration (2 g/L).

7. INDUSTRIAL APPLICATIONS OF EXTREMOZYMES Many extremozymes have immense potential for industrial applications but only few enzymes are utilized for industrial and biotechnological applications due to the lack of effort for their commercialization. Many extremozymes are studied only up to their physicochemical characterization, but large-scale production of many marine enzymes is seldom reported (Muffler & Ulber, 2005). A major drawback for commercialization of extremozymes is deficient in cost-effective large-scale production methods. Extremophiles are major source of many novel commercial enzymes but they require extreme environments for their growth and enzyme production. Boom of biotechnological techniques such as cloning and overexpression of gene of interest in a suitable host can overcome these problems. Cloned host has been grown and overexpressed the respective protein in ambient environmental condition. For continuous supply of enzymes in large quantity, it is essential to carry out intensive research on scale-up and downstream processing. It is pivotal to take efforts for commercialization of existing potential enzymes than screening for an unknown enzyme. Protein engineering is the great research strategy for increasing marine enzymes activity and enhances its tolerance to temperature, pH, organic solvents, or other chemicals such as detergents, etc. Extremozymes have potential applications in food processing, chemical and pharmaceutical industries, bioremediation, and also in biotechnological

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research. More than 60% of world’s enzyme production is attained by protease production. Proteases are used in the textile, detergent, pharmaceutical, and leather industries. Two alkaline serine proteases produced by Bacillus mojavensis showed activity in the presence of several nonionic surfactants as well as solid and liquid commercial detergents and thus they are utilized by detergent industry (Haddar et al., 2009). Lipases are the pivotal enzymes for catalyzing esterification, transesterification, and aminolysis reactions. Several marine microbial lipases are used in paper, cosmetic, food, and detergent industries. Moraxella sp. isolated from Antarctic sea water secreted four cold-adapted lipase enzymes that showed maximum production at very low temperature (Feller, Thiry, Arpigy, Mergeay, & Gerday, 1990). Wang, Chi, Wang, Liu, and Li (2007) screened nine lipase-producing strains such as Candida intermedia, C. parapsilosis, C. rugosa, C. quercitrusa, Y. lipolytica, Rhodotorula mucilaginosa, Lodderomyces elongisporus, A. pullulans, Pichia guilliermondii from 427 yeast strains. Some of these enzymes are able to degrade oil and showed a wide range of potential industrial applications. Novel extracellular phospholipase was obtained from marine Streptomycetes sp. that has optimal activity at pH 8.0 and 45°C (Mo, Kim, & Cho, 2009). K and λ carageenases with molecular masses of 30 and 100 kDa, respectively, were obtained from marine Cytophoga and Pseudoalteromonas, respectively (Hatada et al., 2006; Mo et al., 2009). Cold-adapted xylanases produced by Antarctic microorganisms have been patented and applied in baking industry (Collins et al., 2006; Dutron et al., 2007; Georis, Dauvrin, Hoyoux, Collins, & Feller, 2008). Several marine microbial enzymes have potential to tolerate organic solvents. Esterases and lipases of marine microbial origin revealed high tolerance to organic solvents and these enzymes are consumed by chemical industries for chemical synthesis (Sana et al., 2007). They are utilized for enantioselective product formations as well as for enantiospecific drug molecules synthesis (Rasor & Voss, 2001). Enantioselective alcohol dehydrogenase reported from the hyperthermophilic marine archaeon P. furious is exploited as industrial biocatalysis due to its high organic solvent tolerance (Zhu et al., 2006). Substrate specificity, catalytic properties, and enantioselectivity are extensively studied for this enzyme. It produces enantiomerically pure chiral alcohols by catalyzing the reduction of aryl ketones and its activity is increased by rising temperature without affecting enantioselectivity. Marine microbial enzymes can also be employed for biomedical applications. L-Asparaginase and L-glutaminase were produced by marine bacteria Pseudomonas fluorescens.

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L-Asparaginase has

been used for treatment of children lymphoblastic leukemia and L-glutaminase has antineoplastic activity (Chandrasekaran & Rajeev Kumar, 2003; Kondrat’eva, Dobrynin, & Merkulov, 1978; Schemer & Holcenberg, 1981). Some marine microorganisms have been utilized for bioremediation of industrial and agricultural wastes (Basheer et al., 2011; El-Sersy, Abd-Elnaby, Abou-Elela, Ibrahim, & El-Toukhy, 2010; Kim et al., 2011; Luo, Vrijmoed, & Jones, 2005; Ravindran, Naveenan, & Varatharajan, 2010). Marine enzymes used for degradation of organic wastes produced in seafood farming have many advantages (Trincone, 2012). Crude petroleum and petroleum products are degraded by the marine fungus Geotrichum marinum (Thirumalachar & Narasimhan, 1983). Oil spillage on sea surface is the major pollution in sea which affects the marine ecology. A salt-tolerant petroleum-degrading enzyme from marine microorganisms would be effective in degrading the residual petroleum products.

8. CONCLUSION Marine ecosystem is an immense source of novel enzymes. Marine microbial enzymes have potential to be active at extreme environmental conditions. Some extremozymes have been exploited for biotechnological research, biofuel, and pharmaceutical industries, while many extremozymes reveal potential applicability in environmental management, food processing industries, chemical and pharmaceutical synthesis. With the boom of several high-throughput techniques, it is very easy for the screening of novel enzymes from vast marine ecosystems. Many developments in the field of genetic engineering, accessibility of many biotechnology tools, including a better understanding of protein structure facilitate the development of engineered enzymes with enhanced properties such as higher catalytic activity, more tolerance to organic solvents and chemicals, and higher stability to adverse environments. Furthermore, marine extremophiles may be very good resources for the production of extremozymes which have immense potential in industrial applications.

ACKNOWLEDGMENTS J.S. is grateful to University Grants Commission—Dr. D.S. Kothari postdoctoral fellowship for financial support. Conflict of interest: The authors declare that they have no conflict of interest.

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CHAPTER FOUR

Enzymes From Rare Actinobacterial Strains J. Suriya*, S. Bharathiraja†, P. Manivasagan{, S.-K. Kim{,§,1 *School of Environmental Sciences, Bharathidasan University, Tiruchirappalli, Tamil Nadu, India † CAS in Marine Biology, Annamalai University, Porto Novo, Tamil Nadu, India { Marine Bioprocess Research Center, Pukyong National University, Busan, Republic of Korea § Specialized Graduate School Science & Technology Convergence, Pukyong National University, Busan, Republic of Korea 1 Corresponding author: e-mail addresses: [email protected]; [email protected]

Contents 1. Introduction 2. Rare Actinomycetes and Selective Isolation 3. Various Types of Actinobacterial Enzymes 3.1 Extremophilic Enzymes From Actinobacteria 3.2 Oxidative Enzymes From Actinobacteria 3.3 Lignocellulolytic Enzymes From Actinobacteria 3.4 Other Industrially Important Actinobacterial Enzymes 4. Conclusion Acknowledgments References

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Abstract Actinobacteria constitute rich sources of novel biocatalysts and novel natural products for medical and industrial utilization. Although actinobacteria are potential source of economically important enzymes, the isolation and culturing are somewhat tough because of its extreme habitats. But now-a-days, the rate of discovery of novel compounds producing actinomycetes from soil, freshwater, and marine ecosystem has increased much through the developed culturing and genetic engineering techniques. Actinobacteria are well-known source of their bioactive compounds and they are the promising source of broad range of industrially important enzymes. The bacteria have the capability to degrade a range of pesticides, hydrocarbons, aromatic, and aliphatic compounds (Sambasiva Rao, Tripathy, Mahalaxmi, & Prakasham, 2012). Most of the enzymes are mainly derived from microorganisms because of their easy of growth, minimal nutritional requirements, and low-cost for downstream processing. The focus of this review is about the new, commercially useful enzymes from rare actinobacterial strains. Industrial requirements are now fulfilled by the novel actinobacterial enzymes which assist the effective production. Oxidative enzymes, lignocellulolytic enzymes,

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extremozymes, and clinically useful enzymes are often utilized in many industrial processes because of their ability to catalyze numerous reactions. Novel, extremophilic, oxidative, lignocellulolytic, and industrially important enzymes from rare Actinobacterial population are discussed in this chapter.

1. INTRODUCTION Actinobacteria are primarily saprophytes which remarkably contribute in the turnover of complex biopolymers such as pectin, hemicellulose, keratin, lignocellulose, and chitin. In addition to this, they are also a good resource of antibiotics, enzymes, and other bioactive molecules. Henceforth they are utilized in pharmaceutical and other industries (Stutzenberger & Berdine, 1992). Among the other microorganisms, actinobacteria received much attention due to their production of wide range of biologically active compounds. Chemical reactions are catalyzed by enzymes which can be classified into intracellular and extracellular enzymes based on their origin (Baldrian, 2014). Due to high yield, cost efficiency, and susceptibility to genetic manipulation, microorganisms are mostly preferred for enzyme production. Microbial enzymes have been extensively used in various industries like food, detergent, textile, pharmaceutical industries, and also in the fields of medical therapy, bioorganic chemistry, and molecular biology. Maximum production of commercial enzymes, pharmaceuticals, enzyme inhibitors, and antitumor agents has been obtained from Actinobacteria due its omnipresent nature (Remya & Vijayakumar, 2008). In recent years, numerous research works have been carried out to assess the marine actinobacterial diversity and its enzyme-producing capability in the unexplored marine sediments, hyper saline saltpan, and mangrove. The rare actinomycetes (nonstreptomycetes) are strains of actinomycetes whose isolation rate by culturable methods is much lesser than that of streptomycete strains (Seong, Choi, & Baikm, 2001). Physiology, basic knowledge of the habitats and secondary metabolite diversity of the rare actinomycetes has been gradually increased (Tiwari & Gupta, 2012). Different unexplored environments are targeted for the isolation of rare actinomycetes due to its potential in the production of enormous novel bioactive compounds. A 220 rare actinomycete genera have been isolated from unexplored habitats such as different marine sources (Goodfellow

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et al., 2012), hyper arids (Okoro et al., 2009), soil samples (Ara, Bakir, Hozzein, & Kudo, 2013), Volcanic zones (Lee & Lee, 2011), plant materials (Zhao et al., 2011), extreme saline zones (Jose & Jebakumar, 2013), glaciers (Zhang et al., 2012), and much more. Rare actinomycetes are the good sources of novel enzymes. The isolation of novel enzymes from these rare actinobacteria nourishes the current enzyme discovery programs. Oxidative enzymes are oxidoreductases that catalyze oxidation– reduction reactions. Interest toward the production of actinobacterial oxidative enzymes is increased much over the past 20 years particularly in the fields of lignin degradation and detoxification of organic pollutants, phosphate pesticides, and azo dyes (Pasti, Pometto, Nuti, & Crawford, 1990; Torres, Bustos-Jaimes, & Le Borgne, 2003). Various actinobacterial oxidative enzymes play an important role in morphogenesis or antibiotic production (Endo et al., 2003; Suzuki, Furusho, Higashi, Ohnishi, & Horinouchi, 2006). Lignocellulases are hydrolytic enzymes which are able to degrade tough lignocelluloses such as hemicellulases, cellulases, and lignolytic enzymes (Mtui, 2012) and are also utilized in various applications (Deswal, Sharma, Gupta, & Kuhad, 2012). Lignocelluloses are the abundant renewable biomass on earth (Isikgor & Becer, 2015). Lignocellulolytic enzymes can be produced from diverse types of microbes including fungi and bacteria (de-Souza, 2013). Compared with other bacteria, actinomycetes are an attractive group, being exploited for production of lignocellulases. Lignocellulolytic enzymes, one of the strong compounds delivered by actinomycetes, can be utilized generally as a part of different lignocelluloses based commercial enterprises (Prakash et al., 2013). Cellulases are used in biomethane and bioethanol production (Gupta, Samant, & Sahu, 2012), and also industries like textile industry, detergents industry, pulp and paper industry, animal feed, and food industry (Sukumaran, Singhania, & Pandey, 2005). Hemicellulases are applied in deinking of paper waste, biobleaching, up gradation of feed, fodder, and fibers, clarification of fruit juices, saccharification of hemicelluloses to xylose sugars (Soni & Kango, 2013). A wide range of enzymes useful in biotechnological industries and biomedical fields have been studied from various genus of actinobacteria. While there is essential information available due to the arrival of genomic and proteomic reports of actinobacteria has been constantly screened for the production of novel enzymes like proteases, chitinases, cellulases, xylanases, amylases, and other industrially important enzymes.

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2. RARE ACTINOMYCETES AND SELECTIVE ISOLATION Isolation of rare actinobacterial strains from the environment, culturing them in normal lab condition and maintenance of that isolates under suitable conditions are very hard and difficult which lead to the comparatively low occurrence of rare actinomycetes (Berdy, 2005). In order to overcome this problem, suitable isolation and selection procedures are required (Khanna, Solanki, & Lal, 2011; Qiu, Ruan, & Huang, 2008). Some necessary things have been kept in mind during the isolation of rare actinobacterial strains which include (i) employ appropriate selective media containing macromolecules such as chitin, casein, and humic acid for promotion of growth rate of rare actinomycetes and concurrently reduce contamination by bacterial/fungal colonies (Bredholdt, Fjaervik, Johnsen, & Zotchev, 2008; Cuesta, Garcia-de-la-Fuente, Abad, & Fornes, 2012; Hong et al., 2009; Qiu et al., 2008; Zhang & Zhang, 2011), (ii) by adding different antibiotics for fungal and bacteria to the media will enhance the selection of actinomycetales (Hong et al., 2009; Qiu et al., 2008; Zhang & Zhang, 2011), (iii) Sporulating actinobacterial strains are induced to produce motile spores and it is attained by applying chemoattractants like chloride, xylose, bromide, vanillin, and collidine for spore accumulation from actinoplanes (Hayakawa, 2008), (iv) from the application of different frequency of radiation, selective isolation of various kind of actinomycetes can be done (Bredholdt et al., 2007). SHF (super-high frequency) radiation allow effective isolation of Rhodococcus and Streptosporangium species, EHF (extremely high frequency) and UV radiation applied for efficient isolation of actinomycetes like Nocardiopsis, Nocardia, and Streptosporangium spp. (v) by applying chloramine treatment, Microtetraspora, Herbidospora, Streptosporangium, and Microbispora genera can be selectively isolated (Hong et al., 2009), (vi) for the isolation of marine actinomycetales Salinispora, seawater based media is used (Maldonado et al., 2005). Among numerous actinomycetes species, only 11 rare actinomycetes species producing 50 bioactive compounds altogether were identified in 1970 (Berdy, 2005). Today the number has reached nearly 100 of rare actinomycetes and the number is increasing due to the newly developed genetic and proper isolation techniques (Berdy, 2005; Bredholdt et al., 2007). The ability to produce potential new compounds will certainly make a way for the production of clinically important antibiotics (Anzai et al., 2008; Arumugam et al., 2009; Carlson, Li, Burr, & Sherman, 2009; Hohmann et al., 2009;

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Hong et al., 2009; Lam, 2006; Niu et al., 2007; Pimentel-Elardo et al., 2010; Rahman et al., 2010; Xu et al., 2010). Extremophiles produce extremozymes which have immense potential to work under harsh conditions, which were unsuitable for the normal enzymatic activity. Extremozymes are the promising alternatives for current industrial applications because which has optimal activity and stability under extreme conditions where as rare actinomyces are usually found and active in the common as well as uncommon environments but their isolation prevalence by culture dependent methods is much lesser than that of other actinomycetes strains.

3. VARIOUS TYPES OF ACTINOBACTERIAL ENZYMES Based on source, habitats, uses, and applications, here we divide the actinobacterial enzymes into some divisions for the easy and better understanding.

3.1 Extremophilic Enzymes From Actinobacteria Generally microbes are not restricted to specific environmental conditions. Microbial population can be found in the most diverse conditions, including extremes of pressure, temperature, pH, and salinity. These microorganisms known to be extremophiles, from which we can obtain biocatalysts and these are stable and functional at extreme conditions. Over the past years, industrial application of biocatalysts which can withstand at harsh environment has increased greatly. This is possible by the discovery of novel enzymes from extremophilic microbes from unusual extreme environments. Like other microorganisms, actinobacteria can survive in mesophilic, halophilic, acidophilic, and thermophilic conditions and they have the capability to degrade starch by hydrolysis (Williams et al., 1983). The mesophilic actinomyces genera Nocardia and Streptomyces produce amylases (Mordarski, Wieczorek, & Jaworska, 1970). Studies show that their enzymes are similar to bacillus amylases which are thermolabile. From the literature survey, Streptomyces hygroscopicus (McKillop, Elvin, & Kenten, 1986), S. limosus (Fairbairn, Priest, & Stark, 1986), and S. praecox (Suganuma, Mizukami, Moari, Ohnishi, & Hiromi, 1980) strains are more capable as amylase producers as a result of broad screening methods employed. High concentration of thermostable amylases was obtained from thermophilic actinobacteria, Thermomonospora curvata, T. vulgaris (Stutzenberger & Carnell, 1977; Stutzenberger, Kanno, Tamura, & Suekane, 1978) and from

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a Thermoactinomyces sp. (Obi & Obido, 1984a, 1984b). The compounds are relatively stable and highly active at 60–70°C and also stable at slightly acidic and neutral pH values (Kuo & Hartman, 1967). DyP-type peroxidase enzyme from the thermophilic actinomycete, Thermobifida fusca was isolated by Van Bloois, Torres-Pazmin˜o, Winter, and Fraaije (2010), which show maximum reactivity toward anthraquinone dyes, and sensible activity toward aromatic sulfides, standard peroxidase substrates, and azo dyes. In 2014, Jaouadi et al. isolated humic acid biodegrading peroxidase enzyme from Streptomyces albidoflavus which are highly thermophilic and exhibit higher catalytic efficiency than HRP (Jaouadi et al., 2014). Sutherland, Crawford, and Pometto (1981), when inducing the culture of four thermophilic Streptomyces with benzoic acids, observed high C12O activity. Thermotolerant enzymes obtained from actinobacteria are listed in Table 1. Marine Halophilic Actinobacteria Micromonospora, Streptomyces, and Rhodococcus are isolated from marine environment (Reiss, Ihssen, & Th€ ony-Meyer, 2011). They also isolated other group of actinomycetes which includes Dietzia, Marinophilus, Salinispora, Salinibacterium, Solwaraspora, Aeromicrobium, Gordonia, Microbacterium, Pseudonocardia, Actinomadura, Nocardiopsis, Saccharopolyspora, Nonomuraea, Williamsia, Streptosporangium, and Verrucosispora (Fernandes, da Silveira, Passos, & Zucchi, 2014a; Machczynski, Vijgenboom, Samyn, & Canters, 2004; Reiss et al., 2011). Endophytic Actinobacteria are inhabited at the internal part of plants, which helps in the protection of host plant against insects and various diseases. Generally, these endophytic actinobacteria are Streptomyces sp., Streptoverticillium, Micromonospora, Nocardia, Kitasatospora, Pseudonocardia, Nocardioides, Kibdelosporangium, Actinopolyspora, Brevibacterium, Actinomadura, Table 1 Some Extremozymes from Extremophilic Actinobacteria Enzyme Producers Enzyme Temperature (°C)

pH

Streptomycetes transformant T3-1

Cellulase

50

6.5

Streptomyces rimosus R6-554W

Lipase

50

9.0–10.0

Thermomonospora fusca NTU22

α-Amylase

60

7.0

Thermomonospora fusca

Xylanase

60–80

7.0 (6–8)

Thermoactinomycetes sp. HS682

Protease

70

11.0

Thermomonospora sp.

Cellulase

60–70

6.0

Streptomyces antibiotius

Cellulase

40–55

5–7

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Glycomyces, Plantactinospora, Microbispora, Polymorphospora, Streptosporangium, and Promicromonospora which are found in the plants. Acidophilic Actinobacteria that generally grows at very low acidic pH range from 3.5 to 6.5 is found in acidic forest and mine drainage soil (Fernandes et al., 2014a; Ska´lova´ et al., 2009). Industrially important acidophilic enzymes are being obtained from the Streptomyces sp. and Kitasatospora.

3.2 Oxidative Enzymes From Actinobacteria Laccase, peroxidase, and tyrosinase are the important group of Oxidative enzymes which catalyses oligomerization, depolymerization, and hydroxylation reactions. Laccase and tyrosinase require molecular oxygen for catalyzing their reactions and produce water. Henceforth, these two enzymes are referred as “green enzymes.” Due to these catalyzing properties, oxidative enzymes are utilized in a variety of industrial processes. In the past decades, most oxidative enzymes were obtained from fungal species. But in recent years, it has become clear that these enzymes are well produced by actinobacterial strains, which are considered as an unexplored resource of oxidative enzymes with large number of industrial applications. 3.2.1 Cholesterol Oxidase Cholesterol oxidases (CO) are very useful enzymes for biotechnological applications because of its ability for the detection and conversion of cholesterol. Actinobacterial CO is being isolated from diverse environment and some of the rare actinomycetes like Corynebacterium spp., Streptomyces spp., Rhodococcus rhodochrous, Mycobacterium spp., Rhodococcus erythropolis, and Brevibacterium spp. are the major producers of the enzyme (MacLachlan, Wotherspoon, Ansell, & Brooks, 2000). Ivshina, Grishko, Nogovitsina, Kukina, and Tolstikov (2005) demonstrated the bioconversion of testosterone with the addition of glucose as cooxidant in the presence of the inhibitor 2,2ʹ-dipyridyl by cholesterol oxidase which is isolated from actinobacteria Rhodococcus strains. Now a days CO from actinobacteria are applied in analytical practices, like cholesterol measurement in biological fluids and to quantify the dehydroepiandrosterone sulfate (DHEAS) in cysts of the human mammary gland duct liquids (Donova, 2007). Streptomyces spp. producing cholesterol oxidases are used as a rich source of insecticidal proteins. Though actinobacterial cholesterol oxidase has more industrial applications, these also been concerned as a causative agent for human diseases. Actinobacterial strain Rhodococcus equi act as a primary pathogen for horses and requires cholesterol oxidase for opportunistic infection in humans

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which will cause membrane damage (Kumari & Kanwar, 2012). Brzostek, Dziadek, Rumijowska-Galewicz, Pawelczyk, and Dziadek (2007) confirmed the presence of another cholesterol oxidase which plays a main role in the M. tuberculosis pathogenesis. So from this we can get the idea about developing a treatment for the opportunistic infections by clearly understanding the virulence factors of cholesterol oxidase enzyme. 3.2.2 Peroxidases Peroxidases are a major group of oxidoreductases enzymes which can contain a heme cofactor in their active sites or redox-active selenocysteine or cysteine residues that catalyze the oxidation of substrate molecules by the hydrogen peroxide (H2O2) as the electron acceptor (Van Bloois et al., 2010). Actinobacteria are known to be a rich source of peroxidases, a novel industrially important enzyme, chiefly in a market which is dominated by the plant horseradish peroxidase (HRP) (Le Roes-Hill, Khan, & Burton, 2011; Mercer, Iqbal, Miller, & McCarthy, 1996; Tuncer, Rob, Ball, & Wilson, 1999). This HRP is the major enzyme for the removal of predominant pollutants such as phenols by enzyme-catalyzed polymerization reaction. It can be also used to alter toxic materials into a lesser amount of harmful substances. DyP type of peroxidase enzyme isolated by Van Bloois et al. (2010) from Thermobifida fusca, a thermophilic actinomycete, which showed high reactivity toward anthraquinone dyes. Many actinobacterial peroxidase enzymes involved in many manufacturing processes like computer chips, adhesives, and linings of cans and drums. 3.2.3 Catechol 1,2-Dioxygenase Aromatic compounds are usually broken down by microbes like bacteria. Bacterial populations frequently possess genes code for enzymes catechol 1,2-dioxygenase which are able to degrade toxins into catechol and protocatechuate (Nair, Jayachandran, & Shashidhar, 2006). These enzymes are widely distributed among actinobacterial strains. From the result of molecular analysis of catechol-degrading bacteria, the gene catA that codes for C12O is detected between several actinobacterial strains which include Gordonia, Streptomyces, Rhodococcus, Mycobacterium, and Corynebacterium (El Azhari, Devers-Lamrani, Chatagnier, Rouard, & Martin-Laurent, 2010; Hamzah & Al-Bahama, 1994; Harwood & Parales, 1995; Shen et al., 2009). When induced the cultures with benzoic acid, Sutherland et al. (1981) proved activity C12O in four extremely thermophilic strains of

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Streptomyces. At the same time, C12O activity was seen in Rhodococcus sp. NCIM 2891 when the culture medium is induced with phenol (Nadaf, Zanan, & Wakte, 2011). Silva et al. (2012) successfully isolated C12O and C23O enzymes from the actinomycetal strain Gordonia polyisoprenivorans and checked their activity in the range of polluted environment. Immobilized C120 and C230 enzymes showed higher activity which leads to the greater industrial applications. The thermolabile C12O and C23O enzymes from rare actinomycetes genera take part in the breakdown of contaminants from the polluted agricultural environment, which includes aniline and its derivatives (Kaminski, Janke, Prauser, & Fritsche, 1983) but also the degradation of diesel, biodiesel, dibenzothiophene, and chlorinated benzenes (Field & Sierra-Alvarez, 2008; Silva et al., 2012).

3.2.4 Tyrosinase Tyrosinase is a copper-containing enzyme which usually occurs in plant and animal tissues that catalyzes the production of melanin and other pigments by oxidation from the substrate tyrosine, as in the blackening of a peeled or sliced potato exposed to air. As for now, totally three kind of crystal structured tyrosinases have been studied and one among them was isolated from the actinomycete genera, Streptomyces castaneoglobisporus (Toshio, Suzuki, Asano, Matsuzaki, & Nakamura, 1979). These enzymes are omnipresent in nature and provide a huge number of biological functions (Kurane, Suzuki, & Fukuoka, 1984). In plants, tyrosinases are responsible for the browning of open surfaces present in fruits (Muzzarelli, 1985), but in the case of microbes, the defense mechanism of its DNA against reactive oxygen species and radiation by melanin and also binds with the toxic heavy metals (Cambou & Klibanov, 1984; Chakrabarti, Matai, & Chandra, 1978). Biologically active melanin shows loads of advantages like providing UV radiation protection and antitumor activity (Dadachova et al., 2007; Goncalves & PombeiroSponchiado, 2005; Hung, Sava, Hong, & Huang, 2004; Montefiori & Zhou, 1991). In earlier days fungus are utilized as the major source of tyrosinase enzyme (Sambasiva Rao et al., 2012) but now a days actinobacteria are well-known producers of tyrosinases, particularly many Streptomyces species which produce a melanin-like pigment (Kohashi et al., 2004), and due to this, actinobacterial tyrosinases are increasing widespread (Della-Cioppa, Garger, Holtz, McCulloch, & Sverlow, 1998; Della-Cioppa, Garger, Sverlow, 1998; Matoba, Kumagai, Yamamoto, Yoshitsu, & Sugiyama, 2006).

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3.2.5 L-Aminoacid Oxidase L-amino acid oxidases (L-AAO) are the enzymes belong to oxidoreductases that catalyze the oxidative deamination process of L-amino acids to get ketoacids, hydrogen peroxide, and ammonia (Geueke & Hummel, 2002). These enzymes demonstrate broad substrate ranges and are commonly applied for the resolution of racemic mixtures. An L-amino acid oxidase enzyme isolated from rare actinobacteria Rhodococcus opacus DSM 43250 which exhibit a wide range of substrate that includes the amino acids L-phenylalanine, L-alanine, L-lysine, and L-leucine. This enzyme can be able to resolve a racemic mixture of D,L-phenylalanine and D,L-leucine (Geueke & Hummel, 2002). 3.2.6 Putrescine Oxidase Putrescine is a low molecular weight diamine that belongs to the family of compounds termed biogenic amines. Such amines accumulate in foods and putrescine can act as a marker to detect food spoilage caused by Enterobacteriaceae and Clostridium spp. (Santos, 1996; Shalaby, 1996). Putrescine oxidases (PuOs) are the microbial enzyme which catalyze the oxidative deamination of putrescine into 4-aminobutaral, hydrogen peroxide, and ammonia (Agostinelli et al., 2004). Putrescine Oxidases are isolated from numerous actinobacteria, especially from R. erythropolis and Kocuria rosea (Micrococcus rubens) (Adachi, Yamada, & Ogata, 1966; Van Hellemond, Van Dijk, Heuts, Janssen, & Fraaije, 2008). Thin-layer chromatography, ultraperformance liquid chromatography, and gas chromatography are standard analytical methods used to detect biogenic amines (Kolisis & Thomas, 1987; Ray, 1977). Later newer detection methods like biosensors have now been developed. Immobilized putrescine oxidase from actinobacteria K. rosea with multiwalled carbon nanotubes used as a biosensor for the detection of putrescine present in mammalian plasma, with some interference from cadaverine or histamine, and this sensor does not need any prior sample purification (Kunimoto, Aoyagi, Takeuchi, & Umezawa, 1974). 3.2.7 L-Glutamate Oxidases oxidases are strong substrate-specific amino acid oxidases (Costello, Cisar, Kolenbrander, & Gabriel, 1979). The first actinobacterial L-glutamate oxidase was isolated from Streptomyces violascens (Kamei, Asano, Suzuki, Matsuzaki, & Nakamura, 1983). Glutamate oxidases involved a major role in pharmaceutically relevant chiral intermediates L-glutamate

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synthesis especially glutamate to α-ketoglutarate conversion (Patel, 2001a, 2001b). Some of the oxidative enzymes derived from actinobacteria and its applications are listed in Table 2.

3.3 Lignocellulolytic Enzymes From Actinobacteria Lignocellulose is the most plentiful biomass on earth (Isikgor & Becer, 2015). Tons of lignocellulosic wastes are generated significantly by agricultural, forest, and agroindustrial activities annually and they are present in the readily procurable, renewable feedstock, and economically affordable for various lignocelluloses based applications. Lignocellulolytic enzymes are cellulases, hemicellulases, and lignolytic enzymes which play a key role in the processing of lignocelluloses and these are prerequisite for their consumption in various processes. Lignocellulolytic enzymes are the much potent enzymes predominantly produced by actinomycetes which can be widely utilized in various industries based on lignocelluloses (Prakash et al., 2013). Lignocellulases are hydrolytic enzymes which are used to degrade tough lignocelluloses present in the plant biomass including hemicellulases, cellulases, and lignolytic enzymes (Mtui, 2012). Lignin-degrading enzymes have many applications like the recalcitrant lignocellulosic biomass pretreatment for biofuel production, used in various industries like paper industry, textile industry, cosmetic industry, pharmaceutical industries, food industry, wastewater treatment plants, bioremediation processes, and organic synthesis (Abdel-Hamid, Solbiati, & Cann, 2013). Generally actinomycetes the major and attractive groups of lignocellulose producers from which we can get obtain much amount of desired enzymes (Kumar, Biswas, Soalnki, Kumar, & Tarafdar, 2014; McCarthy, 1987; McCarthy & Williams, 1992; Prakash et al., 2013; Vetrovsky, Steffen, & Baldrian, 2014). Celluloses are the components which are used in many industries such as manufacturing of paper and textile fabric industries and major role in the biofuel production from fermentable glucose as inert packing material (Shokri & Adibkia, 2013). Hemicelluloses posses a wide range of applications over various industries and are also used as dietary fiber because of its nontoxicity and biodegradability (Dhingra, Michael, Rajput, & Patil, 2012). Hemicelluloses act as edible coating agent above the packed foods for their stabilization and also used in paper making because of its adhesive properties. In ice-creams and other foods, xyloglucans and β-glucans are used during their stabilizing, gelling, and thickening processes.

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Table 2 Some Oxidative Enzymes From Actinobacteria Enzyme Producers Enzyme Applications

Streptomyces sp. Cholesterol Produces insecticidal proteins oxidase Rhodococcus equi Cholesterol Membrane damage in host, opportunistic infection oxidase in severally immune-suppressed humans (HIV infection) required COX for their infection in host Nocardia

Cholesterol Serum cholesterol measurement oxidase

Rhodochorus erythropolis

Cholesterol Biocatalysis, oxidations of cyclic allylic, bicyclic, oxidase and trycyclic alcohol to synthesize several ergot alkaloids

Streptomyces natalensis

Cholesterol COX-encoding gene (pimE), COX act as a oxidase signaling protein for the biosynthesis of polyene macrolide pimaricin which produce antifungal antibiotic used in food industry; biosynthesis of polyene macrolid pimracin

Streptomyces virginiae

Cholesterol Cloned cholesterol oxidase choL, fragment from oxidase Streptomyces virginiae is used in the oxidation of diosgein to 4-ene-3-keto steroids

Streptomyces lavendulae

Laccase

Biodegradation of xenobiotic compounds, pulp delignication, textile dye bleaching

M. thermophile

Laccase

Fuel industry

S. coelicolor

Laccase

Degradation of dyes used by textile industry

Rhodococcus ruber

Bioremediation

SilA, Streptomyces ipomoea

Lignin degradation

Streptomyces albus

Peroxidase

Biobleaching of kraft pulps

Thermobifida fusca

Peroxidase

Dye industry

Nacardia alba MSA10

Tyrosinase

Food industry, antimicrobial activity, bioremediation of phenol contaminated waters

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Table 2 Some Oxidative Enzymes From Actinobacteria—cont’d Enzyme Producers Enzyme Applications

Streptomyces violascens

Glutamate oxidases

Key role in synthesis of pharmaceutically relevant chiral intermediates

Kocuria rosea

Putrescine oxidase

Used as a biosensor

Hemicelluloses are good source of xylose sugars, which can be easily fermented to form ethanol fuel (G’ırio et al., 2010). For the hydrolysis of hemicellulose, both enzymatic and chemical methods are being used. While in chemical hydrolysis method, there is a use of expensive chemicals, tough conditions, and have some other limitations too. But lignocellulolytic enzymes play a key role in hydrolysis so this method is suitable for industrial applications. Recently utilization of biomass is performed through biorefinery, with significant utilization of all available components and the amount of waste should be less. Hence lignocellulases perform vital functions in biorefinery processes. The genes from numerous lignocellulolytic actinomycetes have been isolated and successfully expressed in different microbes. Gene code for the enzymes GH1 and GH3 from the actinomycetes C. fimi ATCC 484 were cloned in Escherichia coli showed effective hydrolysis of xylanases and celluloses (Gao & Wakarchuk, 2014). Streptomyces reticuli have cel1 gene coding for the enzyme avicelase which has the capacity to hydrolyze crystalline cellulose more efficiently (Schrempf & Walter, 1995). This particular gene was cloned and overexpressed in other microbes like Bacillus subtilis, E. coli, and Streptomyces spp. and owing to the absence of essential regulatory factors, amount of the enzyme was somewhat low (Walter & Schrempf, 1995). The genes xylI and xylII are isolated from xylose-degrading bacteria Actinomadura sp. strain FC7 are cloned and expressed in Streptomyces  thier et al., 1994). From the actinomycetes Streptomyces sp. S27, lividans (E xylanase gene xylBS27 has been cloned and expressed successfully in Pichia pastoris and the cloned product hydrolyzed xylan to xylobiose effectively (Li, Shi, Yang, et al., 2009). Likewise the laccase gene isolated from Streptomyces coelicolor expressed in Streptomyces lividans which produced large amount of laccase with high purity (Dube, Shareck, Hurtubise, Daneault, & Beauregard, 2008). The thermostable laccase gene REN-7 obtained from Streptomyces lavendulae was cloned in E. coli (Suzuki et al., 2003).

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Gene coding for lignin peroxidase get from Streptomyces viridosporus T7A and this gene is cloned into genetically engineered Streptomyces lividans TK64 which showed better lignocellulose degradation than the normal S. lividans TK64 (Wang, Bleakley, Crawford, Hertel, & Rafii, 1990). 3.3.1 Cellulases Cellulases are mainly produced by microorganism like fungi and bacteria that catalyze the breakdown of cellulose materials into simpler monosaccharides. These enzymes are glycosyl hydrolases which are categorized into some families. Cellulases are generally subdivided into four major classes with regard to the mode of action and substrate specificity. These are exoglucanases, endoglucanases, β-glucosidases, and cellobiohydrolases (del-Pulgar & Saadeddin, 2014; Sadhu & Maiti, 2013). These enzymes are specifically important in hydrolyzing crystalline cellulose (delPulgar & Saadeddin, 2014). Among various actinomycetes, Cellulomonas fimi, Streptomyces thermodiastaticus, Thermomonospora curvata, Streptomyces viridosporus and Streptomyces setonii, Microbispora bispora, and Thermobifida fusca are known to be cellulase producers with high yield (Lynd, Weimer, Van Zyl, & Pretorius, 2002; Wilson, 1992). Thermophilic actinomycete Thermobifida fusca is a spore forming actinobacteria reported to produce cellulose-degrading enzyme (Lykidis, Mavromatis, Ivanova, et al., 2007). Cellulomonas fimi produce free cellulases which does not contain cellulosomes (Christopherson et al., 2013). Likewise, Cellulomonas flavigena, a facultative anaerobe, is also reported as a producer of free cellulases that hydrolyze hemicelluloses and celluloses efficiently. The extracellular cellulases are secreted by means of both sec general secretion system and sec independent twin-arginine translocation (TAT) systems. The actinomycetes T. bifida secrete cellulases by utilizing both secretion systems and another actinomycetes S. coelicolor majorly depends on TAT systems for the export of proteins (Lykidis et al., 2007). Various research reports evidently show higher cellulose degradation capacity of actinomycetales than fungal species. 3.3.2 Hemicellulases Xylan, glucuronoxylan, arabinoxylan, mannan, and xyloglucan are the major abundant components of hemicelluloses. Hemicellulases are generally synthesized combined with cellulases enzymes (del-Pulgar & Saadeddin, 2014; Lynd et al., 2002). Earlier studies have specified the production of cellulase free xylanases from Streptomyces roseiscleroticus (Grabski, Forrester,

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Patel, & Jeffries, 1993; Grabski & Jeffries, 1991) and Saccharomonospora viridis (Roberts & Wan, 1998). For the complete hydrolysis of xylan materials, enzymes like endo-1, 4-β-xylanases, α-L-arabinofuranosidases, β-D-xylosidases, acetyl xylan esterases, α-glucuronidases, and ferulic/ coumaric acid esterases are mainly involved. Mannan is hydrolyzed by synergistic action of β-mannosidases, mannanases, and α-galactosidases by means of cleaving β-1,4 linked mannose from nonreducing ends, hydrolyzing β-1,4-glycosidic bonds internally and removing terminal D-galactosyl residues, respectively (Gilbert, 2010). Cellulose hydrolysis is enhanced by the degradation of mannan and xylan was proven by some studies but they are known to inhibit the activity of cellulase (del-Pulgar & Saadeddin, 2014). Many rare strains of actinobacteria show a very good productivity of hemicellulases for example T. bifida and other actinomycetes reported to produce several xylanases like β-1,4-endoxylanases xyl10A, xyl10B, and xil11A, α-L-arabinofuranosidases, xyloglucanases, xylosidases, α-N-arabinofuranosidases xil43, and β-1,3-glucanases GH81 (del-Pulgar & Saadeddin, 2014; Lykidis et al., 2007). Extracellular endo- and exoxylanases such as β-mannanase, xylan binding domain CBM4, xel74, and mannosidase are primarily synthesized by actinomycetes strain Cellulomonas fimi (Christopherson et al., 2013). Extracellular xylanase production has been also detected in Streptomyces aureofaciens (Jeffrey, Norzaimawati, & Rosnah, 2011), Microbispora siamensis (Boondaeng, Tokuyama, & Kitpreechavanich, 2011), and Streptomyces coelicolor when they are grown on agricultural wastes like sugarcane bagasse, orange peel, pomegranate peel, pineapple peels, etc. (Padmavathi, Thiyagarajan, Ahamed, & Palvannan, 2011). Xylanases are the main hemicellulolytic enzyme, which has been majorly produced from Streptomyces sp. 7b (Bajaj & Singh, 2010), Streptomyces sp. CD3 (Sharma & Bajaj, 2005), Thermomonospora fusca (Ball & McCarthy, 1988; McCarthy, Peace, & Broda, 1985), Streptomyces sp. PC22, x SWU10, Streptomyces sp. MDS, and Streptomyces sp. 234P-16I (Thomas, Joseph, Arumugam, & Pandey, 2013). Cellulomonas flavigena ATCC 482 synthesize an unusual mixture of nearly 19 endoxylanases, combined with GH11,GH10, and GH30 xylanases, β-xylosidase GH43, α-glucuronidase, GH51 α-arabinofuranosidase, mannans GH13 and GH26, and β-glucanase GH81 and GH16. Streptomyces flavogriseus showed good productivity of β1,4-glucan glucanohydrolase (Ishaque & Kluepfel, 1980). By using papyrus, rice straw pulp and cotton stalk pulp, Streptomyces chromofuscus, S. albus, and S. rochei have shown the production of xylanase. Then the obtained xylanase used in the study of bleach effects showed improved brightness in the

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presence of EDTA (Nagieb, Milegy, Faat, & Isis, 2014; Rifaat, Nagieb, & Ahmed, 2005). Streptomyces albus and Streptomyces hygroscopicus have shown effective production of biogas by their own xylanases enzymes using oil cake and straw waste as a substrate (Priya, Stalin, & Selvam, 2012). In a study by Ninawe, Lal, and Kuhad (2006), three strains of Streptomyces namely Streptomyces cyaneus, S. caelestis, and S. tendae were found to be excellent producers of xylanolytic enzyme. Tsujibo et al. (2004) demonstrated the production of acetyl xylan esterases and α-L-arabinofuranosidases enzymes from Streptomyces thermoviolaceus OPC-520. Bhosale, Sukalkar, Uzma, and Kadam (2011) estimated production of xylanases as 326 IU/mL from the actinobacteria Streptomyces rameus and sugarcane bagasse combined with peptone and dextrose are act as substrates. When the strain Streptomyces pseudogriseolus was subjected to UV mutagenesis, the xylanase production was improved (Abdel-Aziz, Talkhan, Fadel, AbouZied, & Abdel-Razik, 2011). Thermoactinomyces like Thermomonospora curvata, Thermomonospora alba, and other strains such as Micromonospora, Microbispora bispora, Nocardia, and Saccharomonospora viridis have shown production of acetylesterases, β-xylosidases, and arabinofuranosidases (Ball & McCarthy, 1988). Xylanase production has also been observed in various actinomycetes by various researchers and they are Microtetraspora flexuosa (Berens, Kaspari, & Klemme, 1996), Streptomyces chattanoogensis CECT 3336 (Lo´pez-Ferna´ndez et al., 1998), Streptomyces chattanoogensis UAH 23 (Ferna´ndez et al., 1995), Thermoactinomyces thalophilus (Kohli, Nigam, Singh, & Chaudhary, 2001), Streptomyces violaceoruber (Khurana, Kapoor, Gupta, & Kuhad, 2007), Streptomyces thermocyanaeviolaceus (Shin et al., 2009), Thermomonospora sp. (Ristroph & Humphrey, 1985), and Streptomyces lividans (Herna´ndez-Coronado et al., 1997; Kluepfel, Shareck, Mondou, & Morosoli, 1986; Morosoli, Bertrand, Mondou, Shareck, & Kluepfel, 1986). Hemicellulolytic mannanase enzyme was produced by Microbispora sp. (Jeffrey, 2008), and the other studies have reported the production of β-xylosidases from S. nitrosporeus, Streptomyces albogriseolus, and Micromonospora melanosporea (Van Zyl, 1985). 3.3.3 Lignolytic Enzymes Lignin degradation is done by means of enzyme complex which predominantly contains three enzymes such as laccases, lignin peroxidases, and manganese peroxidases (Mason et al., 2001; Pla´cido & Capareda, 2015). Laccases are the oxidoreductases that indicate the degradation of polyphenol, the major recalcitrant component present in the lignocellulose (Abdel-Hamid et al., 2013; Madhavi & Lele, 2009) and it requires oxygen as a second

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substrate for the enzymatic action. Microbial laccases, or laccase-like enzymes, are increasingly well-known compounds because of its wide industrial applications. Up to 2014, less than 10 actinobacterial laccases belonging to Streptomyces spp. have been fully characterized (Fernandes et al., 2014a). Compared with other bacterial laccases, actinobacterial laccases have more advantages because they possess suitable characteristics which make them fit for industrial applications with increased thermostability, a wide range of pH and stability under denaturing conditions. Laccase is a multicopper blue oxidase that couples the four electron reduction of oxygen with the oxidation of a broad variety of organic substrates, including polyphenols, phenols, anilines, and even certain inorganic compounds (Fernandes, da Silveira, Passos, & Zucchi, 2014b; Fisher & Fong, 2014; Reiss et al., 2011; Ska´lova´ et al., 2009). Typical laccases consist of three domains, where numerous laccases of actinobacterial strains have only two Cu-binding domains, which indicate the structure of the small laccase (SLAC) from Streptomyces coelicolor (Fernandes et al., 2014a, 2014b; Lu et al., 2013; Machczynski et al., 2004). SilA, Ssl1, and SCLAC are some of the two domain laccases isolated from actinobacterial strains Streptomyces ipomoea, Streptomyces sviceus, and Streptomyces sp. C1, respectively (Gunne & Urlacher, 2012; Lu et al., 2013; Molina-Guijarro et al., 2009). Streptomyces griseus, Streptomyces coelicolor, Streptomyces cyaneus, and Thermobifida fusca are some of the producers of small laccases which are either dimers or trimmers (Chen et al., 2013). The enzymes can be used for industries like textile dying, wine cork making, teeth whitening, and many other diagnostic, environmental, and synthetic purposes. Laccases can also be used in bioremediation. Protein ligand docking can be used to anticipate the putative pollutants that can be degraded by actinobacterial laccase. Search for more actinomycetes with high lignolytic yield, using advanced genetic engineering techniques combined with conventional methods. Fernandes et al. (2014a, 2014b) designed specific primers for detection of laccase-like genes in actinomycetes. Arias et al. (2003) have shown laccase production using soya flour by rare actinobacterial strain Streptomyces cyaneus CECT 3335. Veratryl alcohol oxidation and other lignolytic activities have also been described in Streptomyces viridosporus (Ramachandra, Crawford, & Hertel, 1988; Ruttimann, Seelenfreund, & Vicu˜na, 1987). Streptomyces sp. strain EC-22, strain EC1, Streptomyces badius, Streptomyces cyaneus MT813, Thermomonospora chromogena, Thermomonospora fusca, Thermomonospora mesophila, Amncolata autotirophica, and Micromonospora sp. have shown notable activities

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against lignin related compounds (Ball, Betts, & McCarthy, 1989; Godden, Ball, Helvenstein, McCarthy, & Penninckx, 1992).

3.4 Other Industrially Important Actinobacterial Enzymes 3.4.1 Aminoacylase Aminoacylase catalyzes the hydrolysis of acylated D- or L-amino acids to D- or L-amino acids that act on carbon–nitrogen bonds. Aminoacylases play a major role in pharmaceutical industries and this is why the production of this enzyme receives more increasing interest. D-aminoacylases are unusual in microbes, but Szwajcer, Szewczuk, and Mordarski (1981) discovered the occurrence of aminoacylase from the actinobacteria Micrococcus agilis. Many studies have tested the presence of aminoacylase for example 427 strains of Streptomyces and 16 strains of Streptoverticillium were experimented. Among those enzymes, only four strains of streptomycetes like S. roseiscleroticus, S. olivaceus, S. tuirus, and S. sparsogenes were found to be the producers of the enzyme aminoacylase (Sugie & Suzuki, 1980), especially when the medium was supplemented with inducers like D-phenylglycine, D-leucine, D-valine (Sugie & Suzuki, 1978). S. olivaceus and S. tuirus produced D-aminoacylases purified and characterized in accordance with their substrate specificity and both enzymes showed their potential activity at pH 7.0 thus hydrophobic N-acetyl-D-amino acids can be greatly hydrolyzed than the hydrophilic amino acids. Mycobacterium smegmatis and several strains of Streptoverticillium belong to the rare actinomycetes strains which are reported to produce the enzymes penicillin V and L-aminoacylase extracellularly (Matsumo & Nagai, 1972; Oreshina, Penzikova, Levitov, & Bartoshevich, 1982). L-aminoacylase extracted from Streptoverticillium sp. (Borisov et al., 1984) showed a high hydrolytic activity toward aromatic L-amino acids and N-acetylated aliphatic L-aminoacids (Skvortsova, Galeaev, Nys, Svedas, & Savitskaya, 1984). 3.4.2 Protease Proteases catalyze the cleavage of proteins by hydrolyzing peptide bonds and due to this action these are known to be vital for the survival of organisms. Based on the mechanisms utilized for hydrolytic cleavage these proteases can be divided into four different kind of families and those are serine, threonine, cysteine aspartate, and metalloproteinases. Proteases are extensively used in the food, pharmaceutical, detergent, leather, and textile industries (Fan, Zhu, & Dai, 2001; Mozersky, Marmer, & Dale, 2002). Among the extremophilic sources, thermostable proteases have been reported from

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certain haloalkaliphilic bacteria and actinobacteria (Dodia et al., 2008; Thumar & Singh, 2007). The mesophilic actinobacteria have the ability to produce Streptomyces protease pronase 7 M by S. griseus and fradiase 7 M by S. fradiae. Alkaline proteases from the actinobacteria such as S. albidoflavus, S. nigellus, Nocardiopsis, Thermoactinomyces, and Thermomonospora were well characterized (McCarthy et al., 1985). 3.4.3 Chitinase Chitin is a tough, protective, semitransparent polysaccharides and principal components of crustaceans, fungi, and outer coverings of insects. During the enzymatic hydrolysis, chitin can be effectively hydrolyzed by complex of enzymes which contain chitinase and chitobiase. Then the completely deacylated chitin compound is called as chitosane (Muzzarelli, 1985). Chitinolytic complexes generally found in fungi, bacteria, and particularly in actinobacteria. The chitinase have been isolated from the actinomycetes strains like Amycolatopsis (Streptomyces) orientalis (Tominaga & Tsujisaka, 1976), S. griseus (Berger & Reynolds, 1958), S. antibioticus (Jeuniaux, 1966), and several species of Streptomyces spp. (Beyer & Diekman, 1985; Price & Storck, 1975; Tiunova, Prieva, Feniksova, & Kusnetov, 1976). In industrial scale, chitinases are produced in higher amount when the media is supplied with chitin wastes (Muzzarelli, 1985) and isolated and purified chitinases show higher activity at pH 5.0 although more sensitive to temperature. At the same time, Tsujibo et al. (2000) proved that the thermophilic actinomycetes Streptomyces thermoviolaceus OPC-520 actively produces thermophilic chitinases. Same researcher, Tsujibo et al., in the year of 2003 again reported about the alkaliphilic actinomycete, Nocardiopsis prasina that produced chitinase showing activity against fungi Trichoderma sp. The enzyme chitinase from Streptomyces sp. M-20 also exhibits activity against the fungi Botrytis cinerea (Kim, Yang, & Kim, 2003). Streptomyces hygroscopicus produce chitinase enzymes that degrade the chitin present in the stem rot disease producing fungal Sclerotium rolfsii. Thus it can be used along with organic fertilizers to control the stem rot disease (Pattanapipitpaisal & Kamlandharn, 2012). Like these actinobacteria, many chitinase producing novel actinobacterial strains such as Streptomyces aureofaciens CMUAc130 and Streptomyces hygroscopicus (Prapagdee, Kuekulvong, & Mongkolsuk, 2008; Taechowisan & Lumyong, 2003), Streptomyces sp. DA11 (Han, Yang, Zhang, Miao, & Li, 2009), Streptomyces viridificans (Gupta, Saxena, Chaturvedi, & Virdi, 1995), Streptomyces aureofaciens (Taechowisan & Lumyong, 2003), Streptomyces tendae, S. griseus, S. variabilis, S. endus,

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S. violaceusniger (Gherbawy, Elhariry, Altalhi, & El-Deeb, 2012) exhibited effective antifungal activities. 3.4.4 Lipase Lipases, known as triacylglycerol hydrolases, are involved in the digestion and transport of lipids. They are an important group of enzymes which have huge applications in many industries like food, detergent, dairy, and pharmaceutical industries. Actinobacterial lipases play a key role in commercial purposes. Lipases always utilize lipidic carbon, such as fatty acids, glycerols, oils, and tweens as a substrate with organic nitrogen source. From the actinobacterial strain S. lavendulae lipolytic enzyme cholesterol esterase was isolated (Toshio et al., 1979). Rhodococcus (Nocardia) erythropolis is an actinobacteria producing a lipase of the arylesterase group which is able to hydrolyze phthalate esters in to a free phthalic acid and simple n-alcohols (Kurane et al., 1984). These enzymes are active at pH 8.6 and 42°C. The lipase production is also experimented from several Streptomyces strains and reported (Chakrabarti et al., 1978). 3.4.5 Penicillin Amidase Penicillin amidase are biocatalysts able to hydrolyze penicillins. Penicillin acylases are subdivided into three major groups based on their substrate specificity. The penicillin hydrolysis reaction precedes in an alkaline medium and at lower pH values and is reversible. Penicillin amidase production was reported in actinobacteria was seen majorly and trace amount in moulds and yeasts (Hamilton-Miller, 1966; Vandamme & Voets, 1974). Actinobacteria such as Nocardia, Mycobacterium, and Streptomyces are capable to hydrolyze penicillin (Savidge & Cole, 1975; Vandamme, 1980; Vanderhaeghe, 1975). These enzymes are mostly produced intracellularly. They show higher activity in the pH range 7.0 to 8.0.

4. CONCLUSION Actinobacterial enzymes have wide applications in industries and medical fields. In general actinobacterial enzymes are more stable and more active as compared with the enzymes from other organisms. Actinomycetales are unexplored genera which act as a rich resource for oxidative enzymes, lignocellulolytic enzymes, and other industrially important enzymes with potential for biotechnological application. By knowing the genetics and habitats of actinobacteria one can improve the expression system of the

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actinobacterial strains and thus we can improvise easy access. New genetic engineering techniques are now implemented to get a higher yield of novel enzymes from rare actinobacterial strains. The discovery of stable enzymes from extremophilic organisms results in the production of thermophilic, acidophilic, alkaliphilic, and also halophilic enzymes which posses the major utilization in various industries and increased applications in the production of special chemicals, pharmaceutical intermediates, and so on. Sequencing the metagenomes is a recent trend which will clearly give the idea about the rare microorganisms from extreme habitats. From this review we can clearly state that actinomycetes are an essential source of extremozymes, oxidative enzymes, lignocellulolytic enzymes, and other industrially important enzymes. They represent extensive proportion of the aquatic and soil microflora which are responsible for the degradation of biomass present in nature. Numbers of actinomycetes strains are reported as a potential source of novel enzymes. World with a rapid increasing of human population and exhaustion of numerous natural resources, enzyme technology combined with new biotechnological techniques provide good solution to meet many industrial needs and will open up new room of thoughts to the near future.

ACKNOWLEDGMENTS This book chapter was supported by research funds of Pukyong National University in 2015. J.S. grateful to University Grants Commission—Dr. D.S. Kothari Post-Doctoral Fellowship for financial support. Conflict of Interest: The authors declare that they have no conflict of interest.

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Vetrovsky, T., Steffen, K. T., & Baldrian, P. (2014). Potential of cometabolic transformation of polysaccharides and lignin in lignocelluloses by soil Actinobacteria. PLoS One, 9(2), e89108. Walter, S., & Schrempf, H. (1995). Studies of Streptomyces reticuli cel-1 (cellulase) gene expression in Streptomyces strains, Escherichia coli, and Bacillus subtilis. Applied and Environmental Microbiology, 61(2), 487–494. Wang, Z. M., Bleakley, B. H., Crawford, D. L., Hertel, G., & Rafii, F. (1990). Cloning and expression of a lignin peroxidase gene from Streptomyces viridosporus in Streptomyces lividans. Journal of Biotechnology, 13(2-3), 131–144. Williams, S. T., Goodfellow, M., Alderson, G., Wellington, E. M. H., Sneath, P. H. A., & Sackin, M. J. (1983). Numerical classification of Streptomyces and related genera. Journal of General Microbiology, 129, 1743–1813. Wilson, D. B. (1992). Biochemistry and genetics of actinomycete cellulases. Critical Reviews in Biotechnology, 12(1-2), 45–63. Xu, Y., He, H., Schulz, S., Liu, X., Fusetani, N., Xiong, H., et al. (2010). Potent antifouling compounds produced by marine Streptomyces. Bioresource Technology, 101, 1331–1336. Zhang, D. C., Schumann, P., Redzic, M., Zhou, Y. G., Liu, H. C., Schinner, F., et al. (2012). Alpinus sp. nov., a psychrophilic actinomycete isolated from alpine glacier cryoconite. International Journal of Systematic and Evolutionary Microbiology, 62, 445–450. http://dx.doi.org/10.1099/ijs.0.031047-0. Zhang, J., & Zhang, L. (2011). Improvement of an isolation medium for actinomycetes. Modern Applied Science, 5, 124–127. Zhao, G.-Z., Li, J., Huang, H.-Y., Zhu, W.-Y., Zhao, L.-X., Tang, S.-K., et al. (2011). Pseudonocardia artemisiae sp. nov., isolated from surface-sterilized Artemisia annua L. International Journal of Systematic and Evolutionary Microbiology, 61, 1061–1065.

CHAPTER FIVE

Bioprospects of Microbial Enzymes from MangroveAssociated Fungi and Bacteria K. Saravanakumar*,1, N. Rajendran†,1, K. Kathiresan{,2, J. Chen*,2 *State Key Laboratory of Microbial Metabolism, School of Agriculture and Biology, Shanghai Jiao Tong University, Shanghai, China † Government Arts College, Chidambaram, India { CAS in Marine Biology, Faculty of Marine Sciences, Annamalai University, Parangipettai, India 2 Corresponding authors: e-mail address: [email protected]; [email protected]

Contents 1. Introduction 2. Fungal Enzymes 3. Bacterial Enzymes 4. Other Polysaccharide Enzymes 5. Conclusions and Future Prospects Acknowledgments References

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Abstract Mangroves ecosystem provides the unique habitat for the colonization of fungi and bacteria. Interestingly, the enzymes derived from mangroves associated microorganisms have enormous economic value in industries of agriculture, pulp, paper, medicine, sewage treatments, etc. Microbial enzyme activity is required for the metabolism of plants and animals. In addition, the enzymes are also involved in aquatic animal food cycle and degradation of mangroves detritus. However, the understanding of current status of mangroves associated microorganism-derived enzymes and its application is required to improve the future omics studies. Therefore, this chapter is summarizing the current reports and application on enzymes derived from mangroves associated bacteria and fungi.

1. INTRODUCTION Mangroves are growing in between the land and sea or/and estuarine or/and marine open coastal ecosystems (Kathiresan & Bingham, 2001). This 1

Shared first authorship.

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ecosystem is extremely complex in physicochemical and microbial nature because the estuarine is a connecting point of the marine water and terrestrial river water. Therefore, the wide ranges of physicochemical conditions and the nutrient ranges could be observed according to the seasons and the water intake from the river, where only a few organisms can survive, as facultative or obligate forms. There are two forms of microorganisms reported from mangroves estuarine ecosystems, such as facultative form of microorganisms which arrived from terrestrial region and survived in mangrove ecosystem and obligate forms or true marine organisms which are derived from sea during the high tide (Kohlmeyer, 1974). Some microorganisms can survive in extreme conditions near deep sea volcanoes, even at temperatures over 100°C. Marine microorganisms have unique structure and life habitats (Stach, Maldonado, Ward, Goodfellow, & Bull, 2003) and these unique ecological conditions are providing the isolation of unique microorganisms with novel enzyme systems. Numerous novel enzymes are reported from marine microorganisms compared to terrestrial microorganisms; enzymes have been isolated from microorganisms, plants, and animals; but microorganisms are found to be the important source of enzymes due to their biochemical and genetic diversity (Annison, 1992; Niehaus, Bertoldo, Kahler, & Antranikian, 1999). Enzymes obtained from microorganisms are more virulent and stable than those derived from plants and animals (Bull, Ward, & Goodfellow, 2000; Kin, 2006). The microorganisms in marine environment have the antagonism effect with each other for the better utilization of nutrients from marine system. This action could also induce the significant synthesis of microbial enzymes in coastal environments. Because of the complexity of the marine and coastal environment, there are differences between the enzyme productions in terrestrial and marine organisms. All those enzymes are mainly used as food additives (Bernan, Greenstein, & Maiese, 1997; David & Michel, 1999; Harmsen, Prieur, & Jeanthon, 1997). The microbial enzymes have the significant role in mechanism of marine waste management’s. This reaction prompts to secretion of diversified intra- and extracellular enzyme by marine microorganism or adopted as facultative form. A variety of novel microbial enzymes is reported from marine microorganisms with significant industrial applications (Ghosh, Saha, & Mukherjee, 2005; Haefner, 2003). However, there is need to summarize information about enzymes from microorganisms isolated from marine or estuarine and/or mangroves habitat and this will provide better understanding of microbial enzymes and their applications. Therefore, this chapter aimed to summarize the data about

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the bioprospects of marine or facultative form of fungal- and bacterialderived enzymes.

2. FUNGAL ENZYMES Mangrove ecosystem is one of the rich resources for the coastal fungal isolation and biotechnological applications. Many researchers have shown the importance of the mangroves derived fungi on biotechnological applications and industrial uses (Raghukumar et al., 1994; Saravanakumar, Shanmuga Arasu, & Kathiresan, 2013). Some mangrove-associated fungalderived enzymes and their sources are described in Table 1. Especially the mangroves associated fungal-derived enzymes are applied in paper manufacturing, juice and wine industries, bread making, xylitol production, bioremediation of dye, and metal degradation/removal (Kathiresan et al., 2011; Polizeli et al., 2005; Raghukumar, Muraleedharan, Gaud, & Mishra, 2004). The fungi-derived mangroves are important source for lignocelluloses enzymes (Raghukumar et al., 1994; Saravanakumar & Kathiresan, 2013). Mangrove-associated fungus Aplanochytrium sp., is known to remove malachite green dye and chromium from aqueous solution with the production of enzyme (Gomathi, Saravanakumar, & Kathiresan, 2013). Mangrove-derived Trichoderma harzianum anomorph Hypocrea lixii is known to degrade lead and iron from wastewaters (Saravanakumar & Kathiresan, 2015). Mangrove-derived Trichoderma and yeasts are known to produce bioethanol (Saravanakumar & Kathiresan, 2013, 2014). The mangroveassociated fungi Pestalotiopsis microspora and Aspergillus oryzae are producers of alpha amylase (Joel & Bhimba, 2012). A review related to the ecological role and biotechnological potential of mangrove fungi, described much information about the mangrove fungus and their enzyme production and industrial utilization (Thatoi, Behera, Mishra, & Dutta, 2013). Mangroves fungi such as Cladosporium herbarum, Fusarium moniliforme, Cirrenalia basiminuta, Hyphomycetes, Halophytophthora vesicular isolated from the leaves of Rhizophora apiculata is known to produce the lignin cellulose enzymes (Raghukumar et al., 1994). Schizochytrium aggregatum isolated from mangroves leaves of Sonneratia alba and R. apiculata is reportedly to produce cellulase (Bremer, 1995). In addition, fungal cellulases from mangrove forests have been recently reviewed (Hussain, Altenaiji, & Yousef, 2014). Mangroves associated microorganisms play a relevant role in leaves litter decomposition and conversion of mangroves detritus into protein-rich food for the aquatic animals. During the decomposition of mangrove litters the

Table 1 Microbial Enzymes from Marine Fungi Enzyme Mangroves Associated Fungi

Isolation Source

Reference

α-Amylase

Pestalotiopsis microspora VB5 and Aspergillus oryzae VB6

Joel and Bhimba (2012) Mangroves leaves associates: Rhizophora mucronata, Avicennia officinalis, and A. marina collected from Chidambaram area, India

Amylase

Penicillium citrinum JQ249898

Sediment samples of mangrove forest, Odisha, India

Sahoo, Dhal, and Das (2014)

Lignocellulolytic enzyme

Laetiporus sulphureus

Mangroves, Indian ocean Coast in Dar es Salaam, Tanania

Mtui and Masalu (2008)

Lignocellulolytic enzyme

Pestalotiopsis sp. NCi6

R. stylosa trees on trunks and prop Arfi et al. (2013) roots bark, Mangrove forest, Saint Vincent Bay, Southern province, New Caledonia

Amylase, Protease, Cellulase, Pectinlyase, and Lipase

Aspergillus niger, A. ochraceous, A. flavus, A. luchuensis, A. terreus, Halocyphina villosa, Helicascus kanaloanus, Lignicola longirostris, Rhizopus nigricans, and A. sydowi

Mangrove drift woods, Koraiyar Immaculatejeyasanta, Madhanraj, Patterson, and River, Saradi, Sethukuda, and Xavier Munai of Muthupet Panneerselvam (2011) mangrove forest, India

Laccase, Peroxidase, Caseinase, Cyphellaceous fungus Halocyphina villosa Gelatinase, Nitrate reductase, Lipase, Amylase, Cellulase, Laminarinase

Marine Habitat

Rohrmann and Molitoris (1986)

Amylase, Cellulase, Lipase, Chitinase, and Protease

Mangroves decomposing leaves, Kathiresan et al. (2011) Thraustrochytrids sp., Pichia India salicaria, Geotrichum sp., Pichia fermentans, Cryptococcus dimennae, Trichoderma asperellum, T. aaggerssivum, T. spirale, T. polysporum, Trichosporon sp. Aspergillus sp., Fusarium sp.

Lignin-degrading enzymes

Trematosphaeria mangrovei

Algae and sea grasses samples, Abou Keer, Alexandria

Lytic polysaccharide monooxygenases

Pestalotiopsis sp. NCi6

R. stylosa trees on trunks and prop Patel et al. (2016) roots bark, Mangrove forest, Saint Vincent Bay, Southern province, New Caledonia

Pectolytic enzyme

Foliar fungi Colletotrichum gloeosporioides and Coniella musaiaensis

Mangrove plants

Purkait and Purkayastha (1996)

Cellulase, tyrosinase, lipase, L-glutaminase, and L-asparaginase

Endophytic fungi; Acremonium, Cladosporium, Curvularia, and Saccharomyces

Avicennia officinalis, Ernakulam district, India

Job, Manomi, and Philip (2015)

Endoglucanase, cellobiohydrolase, and β-glucosidase

Hypoxlon oceanicum, Julella avicenniae, Lignincola laevis, Savoryella lignicola, and Trematosphaeria mangrovei

Mangroves

Pointing, Buswell, Gareth Jones, and Vrijmoed (1999)

Xylanase

Staganospora sp.

Mangroves decaying wood, Hong Kong

Luo, Vrijmoed Lilian, and Jones (2005)

Mabrouk, Marti, and Morari (2010)

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nitrogen content of the detritus is increased due to microbial exudates and enzymatic activity. In general, the tannin content in mangrove leaves and bark is high. During the initial period of leaf decomposition, the bacterial count is low corresponding to high tannin content. But the fungal colonies are high when tannin is high (Rajendran & Kathiresan, 2000). Fungi like Aspergilli are capable of producing tannase enzyme to degrade tannin (William, Boominathan, Vasudevan, Gurujeyalakshmi, & Mahadevan, 1986). Venkatesan (1981) isolated 35 cellulolytic fungi from mangrove litters. However, during the decomposition of mangrove litters, significant variation has been observed in microbial activity especially enzymatic breakdown of leaves decomposition.

3. BACTERIAL ENZYMES Mangrove environment is a detritus-based ecosystem consisting of numerous microorganisms especially bacteria. They play an important role in mineralization of organic matters and recycle of nutrients. Bacteria possess enzymes for the breakdown of organic matter. Some mangroveassociated bacterial-derived enzymes and their sources are described in Table 2. Protease is an important enzyme used in the industries with a total sale of 60% in the world markets. It is mainly used in the detergents, leather, and medicine (Kumar, Joo, Koo, Paik, & Chang, 2004; Zhang et al., 2005). The endophytic bacterial genera such as Pantoea, Curtobacterium, and Enterobacter were identified from Brazilian mangroves for their enzyme production. Interestingly, 75% of the isolates showed protease activity and 62% of the isolates showed endoglucanases activity (Castro et al., 2014). Endoglucanase and exogluconase activities were also detected in the bacterial strains collected from mangrove sediments of Brazil (Soares et al., 2013). Alkaline protease from Flavobacterium YS9412-130 and their physical and chemical properties were elucidated with the industrial abstraction technology and purification technique at low temperature (Sun et al., 2001). The maximum production of alkaline protease is achieved from a yeast strain Aureobasidium pullulans (Chi et al., 2007). Alkaline protease derived from shipworm bacteria are used as cleaning additive (Greene, Grifin, & Cottal, 1996). Haddar et al. (2009) isolated alkaline protease from a bacteria Bacillus mojavensis A21 and purified two detergent-stable alkaline serine proteases. Alkaline phosphatase gene expression has been reported from Pyrococcus abyssi (Zappa et al., 2001).

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Table 2 Enzymes Isolated from Mangrove-Associated Bacteria MangroveAssociated Bacteria Isolation Source References Enzyme

Alkaline protease

Flavobacterium YS9412-130

Yellow Sea

Sun et al. (2001)

Alkaline protease

Aureobasidium pullulans

Marine Environment

Chi, Ma, Wang, and Li (2007)

Alkaline protease

Bacillus mojavensis A21

Protease

Castro et al. (2014) Endophytic Pantoea, Curtobacterium, and bacteria from Brazilian Enterobacter mangrove plants

Lipases

Moraxella species

Amylase

Bacillus sp.

Decomposing Kathiresan et al. (2011) mangrove leaves

Amylase, esterase, and endoglucanase

Bacillus sp.

Brazilian Castro et al. (2014) mangrove plants

α-Amylase

Streptomyces species Marine Environment

Chitinase or chitosanase

Flavobacterium, Penicillium, Pseudomonas, Enterobacter

Marine Environment

Fukamizo (2000), Monzingo, Marcotte, Hart, and Robertus (1996), and Xia, Liu, and Liu (2008)

Chitinase

Vibrio sp., Aeromonas hydrophila, Listonella anquillarum

Marine Environment

Osama and Koga (1995)

Chitinase A, Chitinase B, Chitinase C

Alteromonas sp. strain 0–7

Marine Environment

Tsujibo et al. (2002)

Haddar et al. (2009)

Feller, Thiry, Arpigny, Mergeay, and Gerday (1990)

Chakraborty, Khopade, Kokare, Mahadika, and Chopade (2009)

Continued

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Table 2 Enzymes Isolated from Mangrove-Associated Bacteria—cont’d MangroveAssociated Bacteria Isolation Source References Enzyme

Agarase

Gram-negative bacteria

Mangroves sediment of Andaman

Shome and Shome (2001)

Agarase

Bacillus, Vibrio, Alteromonas, Cytophaga, Pseudomonas

–

Aoki, Araki, and Kitamikado (1990), Leon, Quintana, Peruzzo, and Slebe (1992), and Hosoda, Sakai, and Kanazawa (2003)

–

Sarwar, Matoyoshi, and Oda (1987)

k-Carrageenase Cytophaga species Cellulase

– Cytophaga, Cellulomonas, Vibrio, Clostridium

Cellulase

Micrococcus, Bacillus, Pseudomonas, Xanthomonas, and Brucella

Mangrove environment

Behera, Parida, Dutta, and Thatoi (2014)

Cellulase

Bacillus cereus JD0404

Mangrove sediment of Thailand

Chantarasiri (2015)

L-Asparaginase

Bacterium

Mangroves of Andaman

Shome and Shome (2001)

Mangrove sediments of Brazil

Soares et al. (2013)

Endoglucanase Bacterial strains and exogluconase

Bras, Cerqueira, Fernandes, and Ramos (2006)

Lipase has various applications, especially in the digestion of fat and oil for the release of free fatty acids and glycerols (Babu, Pramod, & George, 2008). The relevance of lipases has been evident by industrial applications such as paper, cosmetics, food, and beverages (Chi et al., 2009; Kobayashi, Nakajima, & Inoue, 2002), and also used in fish processing (Kojima & Shimizu, 2003). Europe and Japan are the enzyme detergent marketers and their shares reached 90% and 80%, respectively. David (1935) and Kirsh (1935) first discovered microbial lipase expressions and

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their valuable as biocatalysts was described (Amare, Thomas, Steffen, & Per, 2003; Davidson, 2006; Pierre, Jean, Jean, & Michael, 2000). Feller et al. (1990) isolated lipases secreted by Moraxella sp. and other nine lipase producing strains (Wang, Chi, Wang, Liu, & Li, 2007). Amylases are enzymes that break the complex sugars into simple sugar. Due to advancement of marine science and biotechnological research, a large number of microorganisms in the marine habitats are known to produce amylase. A novel α-amylase from marine actinobacteria Streptomyces sp. was isolated (Chakraborty et al., 2009). Bacillus species collected from decomposing mangrove leaves exhibited maximum amylase activity and minimum or no activity for cellulose (Kathiresan et al., 2011). Chitin is a biopolymer widely distributed in nature after cellulose. The chitin and chitosan have similar chemical properties and they have received attention in recent years (Ngo, Kim, & Kim, 2006; Ngo, Qian, Je, Kim, & Kim, 2008). There is a large amount of chitin available in the seawater due to continuous shedding of marine zooplankton (Jeuniaux & Voss-Foucart, 1991). Researchers found a wide variety of microorganisms including Aspergillus, Pseudomonas, Chromobacterium, Clostridium, Flavobacterium, Penicillium, Rhizopus, Enterobacter, Klebsiella, and Streptomycetes that can produce chitinase or chitosanase (Fukamizo, 2000; Monzingo et al., 1996; Xia et al., 2008). Chitinoglastic bacteria and the chitinase activity in the alimentary tract of the lake animals were examined. Chitinase from six marine bacterial species, Vibrio fluvialis, V. parahaemolyticus, V. mimicus, V. alginolyticus, Listonella anguillarum, and Aeromonas hydrophila was reported (Osama & Koga, 1995). Alteromonas sp. strain 0–7 secretes chitinase A (ChiA), chitinase B (ChiB), chitinase C (ChiC) in the presence of chitin (Tsujibo et al., 2002). Agar is a highly heterogeneous polysaccharide with a wide range of applications in the food and beverage industries and also used as moisturizing cosmetic additives (Oren, 2004; Parro & Mellado, 1994; Rasmussen & Morrissey, 2007). Microbial source of agarase enzyme isolated from mangroves of Andaman (Shome & Shome, 2001). Microorganisms such as Cytophaga, Bacillus, Vibrio, Pseudomonas, Alteromonas, Pseudoalteromonas, and Streptomyces are producers of agarase (Aoki et al., 1990; Hosoda et al., 2003; Leon et al., 1992). Researchers have cloned and sequenced several agarase genes, for instance, the Streptomyces agarase gene (dagA), Pseudomonas agarase gene (agrA), and Vibrio agarase gene (agrA) (Belas, 1989; Buttner, Fearnleiy, & Bibb, 1987). Red seaweeds are abundantly present in the coastal environments and are potential source of sulfated polysaccharides especially carrageenan and

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carrageenin. These polysaccharides are mainly used for food and related industries, pharmaceutical, cosmetic industries, and also used as a coagulant, adhesive, and emulsifier (Roberts et al., 2007). Some researchers have found polysaccharides degrading enzymes from bacteria. Sarwar et al. (1987) obtained extracellular k-carrageenase enzyme with molecular weight of 10 and 30 kDa from a bacterium Cytophaga able to degrade the carrageenan. Cellulose is the most available polysaccharide consisting of linear chain of several glucose units. It is used for biotextile industries and processing of cotton and linen products. Cellulases degrade processing waste of seaweeds and can be used as biofertilizer. The breakdown of cellulose through a hydrolysis reaction into smaller saccharides called cellodextrin (Bras et al., 2006). The bacterial strains belonging to the genera Micrococcus, Bacillus, Pseudomonas, Xanthomonas, and Brucella produced the cellulose in the mangrove environment (Behera et al., 2014) and also Cytophaga, Cellulomonas, Vibrio, Clostridium, and Streptomyces produce cellulose. The bacterial strain Bacillus cereus JD0404 isolated from the mangrove swamp sediment was characterized for its cellulolytic activity (Chantarasiri, 2015). Xylanase can hydrolyse polysaccharides of economic value and can depolymerize the plant cell component xylan (Maki, Leung, & Qin, 2009). Bacteria, fungi, yeast, and some marine algae produce xylanases that can contribute to beverage clarification by degrading some polysaccharides in juice or beer (Doi, 2008; Klemm, Heublein, Fink, & Bohn, 2005; Nishiyama, Langan, & Chanzy, 2002). Several marine isolates showed alkaline xylanase activity (Raghukumar et al., 2004).

4. OTHER POLYSACCHARIDE ENZYMES Furukawa, Fujikawa, Koga, and Ide (1992) purified fucoidanase, generated from marine Vibrio and obtained three types of enzymes, which can hydrolyze the substrates to small molecule-oligosaccharides. Yahata, Watanabe, Nakamura, Kamimiya, and Tanaka (1990) isolated a new glucanase from the marine Bacillus bacterium. Araki and Kitamikado (1981) reported that 117 marine bacterial strains belonging to 8 genera produce β-mannanaese. Mahajan et al. (2012) isolated a bacterium, Bacillus subtilis ICTE-1 from marine niche that produces fibrinolytic enzyme. Similarly, Pseudomonas species also produce fibrinolytic enzyme (Liu et al., 2010). This enzyme is mainly used in the cardiovascular ailments. The enzyme isolated from the sulfate-reducing bacterium strain WN specifically catalyzes the demethylation of dimethylsulfoniopropionate (Jansen & Hansen, 2000).

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Martinez, Smith, Steward, and Azam (1996) screened 44 marine isolates and 8 showed the enzyme activity, measured as the hydrolysis of fluorogenic substrates. Taylor and Summer (1987) studied the histamine formation in fish during bacterial spoilage due to the decarboxylation of the amino acid histidine by bacteria with the enzyme histidine decarboxylase. A study of arysulfatase activity and arysulfatase-producing bacteria in sediments collected from marine, estuarine, and mangrove biotopes revealed that from the three biotopes investigated, the mangrove area exhibited maximum activity. L-Asparaginase enzymes were assayed from bacterial strains collected from mangroves of Andamans. Out of 200 isolates, 108 synthesized L-asparaginase (Shome & Shome, 2001). Topoisomerases are key enzymes for adaptation to high temperatures in thermophilic organisms and they were isolated from Thermotoga maritime showing a very efficient action as zinc binding material (Viard, Lamour, Duguet, & Bouthier de la Tour, 2001). Ramaley and Hudock (1973) purified a NADP-specific isocitrate dehydrogenase from thermophilic gram-negative bacterium Thermus aquaticus YT-1 and from B. subtilis-168. The enzyme isolated from the thermophilic organisms was much more thermostable than other organisms. Baumann and Baumann (1973) determined the pattern of allosteric regulation of aspartokinase activity in Beneckea and in the marine luminous bacteria. The results indicated that these organisms have at least three isofunctional aspartokinases of which the first is inhibited by L-threonine, the second one is inhibited by L-lysine, and the third one is unaffected by either L-threonine or L-lysine. A bacteriolytic enzyme was extracellularly produced when Chaetomium globosum, a marine isolate cultivated in a medium with an increased MgCl2 content (Imada, Igarasi, Nakahama, & Isono, 1973). The marine bacterium Pseudoalteromonas atlantica is a producer of cellular and extracellular enzymes (Hoffman & Decho, 2000) and the microbial ectoenzyme activity was studied in an aquatic environment (Ammerman & Glover, 2000). An aldehyde oxidoreductase from Desulfovibrio desulfuricans ATCC27774 was isolated (Susana, Melo, Carita, Teixeira, & Saraiva, 2007).

5. CONCLUSIONS AND FUTURE PROSPECTS Mangroves associated fungal and bacterial enzymes have enormous applications. They are importantly involved in the life cycle of aquatic fish and feed preservation. The microbial enzymes have significant industrial values in agriculture, biofuels, paper industries, bakery, and bioremediation

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of wastewater. Furthermore, the extended studies on the proteomics and transcriptomics mangroves associated microbial enzymes could provide some crucial information about the mangroves associated fungal and bacterial genome on enzyme production and aquatic life cycle in mangroves habitats.

ACKNOWLEDGMENTS The authors are thankful to the authorities of Shanghai Jiao Tong University (K.S. and J.C.), The Principal, Government Arts College (N.R.), and the authorities of Annamalai University (K.K.) for all support. We thank the financial support from the Special Project of Basic Work for Science and Technology (2014FY120900), the key project of the Basic Research of Shanghai Municipal Science and Technology Commission (12JC1404600), the special fund of Modern Agricultural Industry Technology System (CARS-02), and Project 948(2011-G4), Ministry of Agriculture, China.

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CHAPTER SIX

Mechanism and Aquaculture Application of Teleost Enzymes Adapted at Low Temperature C.-L. Wu, B.-Y. Li, J.-L. Wu1, C.-F. Hui1 Institute of Cellular and Organismic Biology, Academia Sinica, Taipei, Taiwan 1 Corresponding authors: e-mail address: [email protected]; [email protected]

Contents 1. Introduction 2. Lactate Dehydrogenase 3. Creatine Kinase 4. Fatty Acids Desaturase 5. Conclusions Acknowledgment References

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Abstract Environment temperature highly influences the physiological condition of poikilothermic teleost. There are different physiological and biochemical responses between fish in different habitats. In order to take profit of fish adapted to different temperatures, some important enzymes have been isolated, assayed, and analyzed. Enzyme expression patterns and properties were evaluated in lactate dehydrogenase, creatine kinase, and stearoyl-CoA desaturase studies. In this chapter, we try to identify the mechanisms of enzyme activity at low temperature by comparing different studies on enzyme kinetics and regulation. The protein–protein interaction between monomers, protein–solvent interaction, and protein substrate correlation are discussed. Studying fish enzymes could accumulate the understanding of marine organism’s enzyme function during adaptation in different temperature zones. Based on these mechanisms, the application of cold-adapted enzymes in aquaculture system is illustrated. Furthermore, this information may create a possible explanation of fish physiological and biochemical evolution route and construct an appropriate strategy to overcome the climate change.

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1. INTRODUCTION Fish, as a poikilothermal animal, changes physiological responses as ambient temperature changes. The ambient temperature is changed from 35°C to 0°C in marine sphere. The metabolism of fish in different temperatures is controlled by more than one metabolic profile when environment temperature changes more than 16°C (Guderley, 1990). Since 1970s, there are chapters discussing fish enzymes and how they may contribute to physiological changes in extreme temperature. Mostly, the cold-adapted mechanism of polar fish and cold water fish has been revealed. Antifreeze glycoprotein of winter flounder (Hew, Liunardo, & Fletcher, 1978) and hemoglobin of icefish (Douglas, Chapman, & Hemmingsen, 1973) have been thought to be an important phenomenon for fish to be adapted to low temperature. On the other hand, biochemical analysis of enzymes in energy metabolism, nucleotide synthesis, proteolysis, and homeostasis contribute to the understanding of molecular and physicochemical mechanism of cold adaptation (Guderley & St-Pierre, 2002; Hazel, 1984; Johnston & Dunn, 1987; Pikaard, 2006). As long as the techniques progressive of analyzing protein structure, structure-base studies of cold-adapted enzymes have bloomed (Marshall, 1997). Function of enzymes adapted to different temperatures could vary by fine structure adjustment. By comparing the homologs of different species, kinetic and thermodynamic properties of psychrophilic, mesophilic, and thermophilic enzymes are clarified (Lonhienne, Gerday, & Feller, 2000). This chapter is focusing on the mechanism of fish cold adaptation and discusses the physiological change of fish, depending on metabolic enzymes. Temperature is a “shape force” in protein evolution of poikilothermal (Somero & Low, 1976). Since hot aquatic environment is unusual in fish living sphere, low temperature is a major factor of fish evolution (Barrett, 2001; Franklin, 1998; Gillis & Tibbits, 2002; Johnston, 2003; Petricorena & Somero, 2007; Verde, Giordano, & Di Prisco, 2008). Protein dynamics across an entire enzyme can play a role in adaptation to differing physiological conditions (Jayasundara, Tomanek, Dowd, & Somero, 2015; Klinman & Kohen, 2014). Biochemical adjustments have minimized the effect of temperature on the catalytic processes of organisms living at widely different temperatures (Guderley, 1990). However, there are few review manuscripts on fish enzymes adaptation at low temperature (Marshall, 1997). Some of important metabolic-related enzymes are discussed in this review. In a further survey, there are couples of enzymes’ structures and

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function revealed since 1997. The relationship between structure, function, and evolution of some enzymes at low temperature are discussed. Since energy supplement is the most important criteria of cold adaptation, energy generation enzymes (Jayasundara et al., 2015; Yancey & Somero, 1978), lactate dehydrogenase (LDH), and creatine kinase (CK) are discussed first. An important membrane fluidity correlated enzyme, stearoyl-CoA desaturase (SCD), is also introduced. Some examples of aquaculture species are also introduced. This may constitute a new approach to increase production of aquaculture during climate change. Cold-adapted enzymes contribute in the secondary response which majorly affects metabolism.

2. LACTATE DEHYDROGENASE Lactate dehydrogenase (LDH, EC 1.1.1.27) catalyzes the interconversion of pyruvate and lactate with concomitant interconversion of NADH and NAD+ (Fig. 1). Muscle form LDH (M4-LDH) homologs were firstly considered in different temperature-adapted species with different activity in 1978 (Yancey & Somero, 1978). This enzyme has been found effective in short-term and long-term temperature driven evolution. At the first, expression form of LDH has been found to be changed in different temperatures from muscle form (high temperature and high anaerobic metabolism) to a heart form (low temperature and high aerobic metabolism) (Somero, 1973). After analyzing the activity of LDH of rainbow trout at different temperatures, a relatively higher activity at 10°C has been found (Sauer & Haider, 1977). In order to rule out the pH influence on enzyme activity, muscle intracellular buffering capacity has been studied (Castellini & Somero, 1981). This study indicated that intracellular buffering capacity of locomotory muscles which was functional under burst locomotion and prolonger, low-level anaerobic condition was necessary. Circulatory perfusion in locomotory muscles was inadequate to rapidly remove the acidic end-products such as lactic acid. In this aspect, white skeletal muscle of teleost had to solve acid–base imbalance by a common biochemical strategy.

Fig. 1 LDH catalyzes pyruvate to lactate companied by NADH to NAD+, and the reverse reaction.

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To reveal the possible mechanism of M4-LDH function in different temperatures, peptides sequences of LDH of four different Sphyraena species, S. argentea, S. lucasana, S. idiastes, and S. ensis (Fig. 2A), were cloned to understand their kinetic properties (Graves & Somero, 1982). Different species which adapted to different environmental temperatures tended to offset temperature effect on metabolic reaction catalyzed by individual enzymes. Three LDH allozymes were highly similar in kcat value between species with different body temperature. Rate compensatory differences in catalytic efficiency have been found sufficient in minor body temperature differences (5–8°C). That is, kinetic differences among distinct enzymes reflected adaptation to different habitat temperatures. In the other family, Sciaenidae, Km of LDH increased in the species adapted at higher temperature (Coppes & Somero, 1990). Consequently, protein structures can provide explanation for LDH thermodynamic status (Low & Somero, 1975). Free energy (ΔG#) and enzyme volume (ΔV#) were changed by interaction of solute and side-chain of amino acids in LDH. In order to realize the molecular conformation of LDH adapted to different temperatures, enzyme of different species of teleost living in different environments were cloned and analyzed. A4-LDH isolated from Antarctic ( 1.86 to 1°C) and South American (4 to 10°C) notothenioid teleosts, demonstrate that (i) the divergent evolutionary thermal histories of the Antarctic and South American species have led to temperature-adaptive changes in A4-LDH kinetics but not in resistance to heat inactivation; (ii) comparisons among notothenioid A4-LDH orthologs indicated that only minor differences in primary structure outside the active site were necessary for modification of kinetic properties; (iii) the coldadapted kinetic properties of notothenioid A4-LDHs appeared to have arisen through changes in conformational flexibility in areas of the molecule that controlled structural movements known to be rate-limiting for catalysis; (iv) these findings suggested a model of enzymatic adaptation to temperature in which cold-adapted orthologs possessed higher conformational entropy, that is, occupied a broader distribution of conformational states at a given temperature than do warm-adapted orthologs (Fields & Somero, 1998). By comparing primary sequence of different teleost A4-LDH, the hot spot of variation located in residues 5–50, 171–189, 212–232, 263–265, 309–316 (Fig. 2B). As the active site of A4-LDH were His193 and Arg169, most variation sites located around the reaction pocket except N-terminal (residues 5–50) arm. The quaternary structure of A4-LDH is shown in Fig. 3. Even primary sequence of A4-LDH could not fully

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Fig. 2 (A) Peptides sequences alignment of different species of M form LDHs. Dogfish (Squalus acanthias); longjaw mudsucker (gillichthys mirabilis); shortjaw mudsucker (Gillichthys seta); blackeye goby (Rhinogobiops nicholsii); pelican barracudas (Sphyraena idiastes); Lucas Barracuda (Sphyraena lucasana); Pacific barracuda (Sphyraena argentea). (B) Molecular modeling of dogfish M-LDH monomer. The most important variation regions to regulate enzyme activity at low temperature were labeled. Arg 169 and His 193 were the active site of M-LDH.

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Fig. 3 Quaternary structure of M4-LDH. Five residues (Lys 6, His 10, Trp 20, His 156, and His 268), which were correlated to subunits interaction at low temperature, of a subunit (green) were labeled.

represent the cold adaptation of some species, such as gobiid fish (Family Gobiidae) (Fields & Somero, 1997). This enzyme could fold into a number of conformers with different stabilities and functional properties. The A4LDH of Gillichthys seta furnishes evidence that such conformers might provide an important mechanism for adaptation of proteins to temperature. In barracuda, evolutionary adaptation of proteins to temperature might be achieved by minor changes in sequence at locations outside of active sites, and these changes might independently affect kinetic properties and thermal stabilities. Structural analysis showed that the residues responsible for temperature adaptation of LDH-As did not reside in the loop region. The differences in the Km of pyruvate between the LDH-As of S. argentea and S. lucasana must involve effects at sites 8 plus 61 and/or 68 (Holland, McFall-Ngai, & Somero, 1997). The crystal structures of these LDHs have been solved. When compared at the structural level, hyperthermophilic, mesophilic, and psychrophilic LDHs have revealed that temperature adaptation was caused by a few amino acid substitutions that were localized in critical regions of the

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enzyme. These substitutions, each having accumulating effect, played a role in either the conformational stability or the local flexibility or in both. From hot- to cold-adapted LDHs studies, the various substitutions have decreased the number of ion pairs, reduced the size of ionic networks, created unfavorable interactions involving charged residues and induced strong local disorder. The analysis of the LDHs adapted to extreme temperatures exhibited how evolutionary processes shifted the subtle balance between overall stability and flexibility of an enzyme (Coquelle, Fioravanti, Weik, Vellieux, & Madern, 2007). A localized increase in conformational entropy was involved in cold adaptation (Fields & Houseman, 2004). In DrLDH, the tryptophan residue at position 79 abolishes the ionic network which was present in TtLDH and contributed to the presence of two negative charges. Six different amino acid residues (Lys 6, His 10, His 20, His 156, His 268, Fig. 3) which were correlated to temperature adaptation have been identified in LDHs of the three hagfishes. These residues were all hydrophilic residues on the molecular surface which might form salt bridges intermolecules and increase enzyme stability. The arrangements of the 22 N-terminal residues were different between Entosphenus japonicus and the three hagfishes. This was thought to be the part where the subunits combine to form a tetramer to change the function of the isozyme. These differences might reflect the tolerance to high pressure and low temperature of the LDHs from the three hagfishes at different habitat depths (Nishiguchi, 2008). To sum up, primary sequences of different LDHs have shown few different residues between related species that live in different temperatures. Fine study on molecular structure of LDH is required to get a universal conclusion to explain the thermodynamic and kinetic properties of LDH adapted at low temperature. However, it is reasonable to say that peptides sequences provide a clue to verify the thermostability of LDH. This is an important criterion for enzyme to stay at different temperature. Also, the secondary structure of LDHs is not changed a lot to present a conclusive explanation. Since LDH acted as a tetrameric enzyme, tertiary structure and quaternary structure variations may control their activity. Most of LDH peptide sequence differences regulate intramolecular and intersubunit interaction to modify their substrates binding affinity and orientation. In this way, activation energy is decreased and enthalpy is increased in enzyme which acted at low temperature. The studies on LDH illustrate that thermostability is correlated to the flexibility of enzyme and regulated its activity at low temperature.

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Fig. 4 Fish growth rate is proportional correlated to LDH activity. Adapted from Mathers, E.M., Houlihan, D.F., & Cunningham, M.J. (1992). Nucleic acid concentrations and enzyme activities as correlates of growth rate of the saithe Pollachius virens: Growth-rate estimates of open-sea fish, Marine Biology 112 (3), 363–369; Jayasundara, N., Tomanek, L., Dowd, W. W., & Somero, G.N. (2015). Proteomic analysis of cardiac response to thermal acclimation in the eurythermal goby fish Gillichthys mirabilis. Journal of Experimental Biology, 218(Pt. 9), 1359–1372.

This increase in LDH activity is able to increase anaerobic component of metabolism during exposure of sea bream (Sparus aurata) to low temperature (Kyprianou et al., 2010). LDH activity was significantly and consistently correlated with the Atlantic cod (Gadus morhua L.) weight and length (Jordan et al., 2005). Since some of amino acids substitutions may promote adaptation of fish (Fields & Houseman, 2004), increasing the levels of LDH in cold-adapted fishes may be attributed to molecular flexibility generated by modifications in structure of enzyme to maintain an appropriate catalytic rate during cold adaptation (Gopakumar & Santhosi, 2009). LDH activity was proportional to growth rate of fish (Fig. 4) (Jayasundara et al., 2015). Traditional gene selection of cold-adapted fish with specific LDH gene structure and expression levels has been evaluated in these studies.

3. CREATINE KINASE Creatine kinase (CK, EC 2.7.3.2) is an enzyme that transfers energetic phosphate group between adenosine nucleotide and creatine (Fig. 5). It is an enzyme which maintains energy homeostasis of cell. The enzyme has been found highly expression at low temperature acclimated at low temperature (Gosselin-Rey & Gerday, 1970). There were four isoforms of CK, including muscle form (M form) and brain form (B form) (Sun, Hui, & Wu, 1998).

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Fig. 5 CK catalyzes creatine to phosphocreatine companied by ATP to ADP, and the reverse reaction.

Muscle form CK was expressed in muscle cytosol and named M-CK. Most mammal have one gene encoded M-CK, however, teleost, including common carp, have more than one subisoform M-CK. There were three M-CK genes in common carp named M1-, M2-, and M3-CK (Gosselin-Rey & Gerday, 1970). Due to M-CK active in dimmer form, as the tetrameric LDH, different combination of isoforms were supposed to be a major effect of enzyme activity and distribution. A study on different subisoform M-CK distribution shows that there were no differences of three subisoforms expression and cellular distribution. At the same time, M-CK expression pattern was not changed during different temperatures treated Atlantic cod (Gadus morhua L.) (Hall, Cole, & Johnston, 2003). They might be different in coupling pattern in different temperatures (Sun, Liu, Hui, & Wu, 2002). In this study, the interaction between protein dimerization area was discussed, a salt bridge between molecules was supposed to be the important factor of M3-CK coupling. Under the assumption, a transgenic M3-CK zebrafish had been generated and it can maintain its swimming ability at low temperature either triggered by CMV (Wu, Lin, et al., 2011) or myosin light chain promoter (Wang, Tan, Jiao, You, & Zhang, 2014). However, M3-CK degradation was also enhanced at low temperature (McLean et al., 2007) that means some other subisoform was responded for M-CK activity at low temperature. M1-CK was found high activity than M3-CK at low temperature and high pH. The activation energy (Ea) of M1-CK was lower than M2- and M3-CK at low temperature based on the kinetic analysis of different M-CKs (Wu et al., 2008). By comparing homologous enzymes from poikilothermal and homeothermal species, their substrates affinities were similar in their surviving temperature, respectively (Ciardiello, Camardella, Carratore, & Di Prisco, 2000; Fields, Wahlstrand, & Somero, 2001). Also, the apparent Km of Chaenocephalus aceratus myotomal CK for ADP was very similar to that of MMCK enzymes from other vertebrates (Cantwell et al., 2001; Nihei, Noda, & Morales, 1961). However, the apparent Km of C. aceratus MMCK for PCr was 5- to 20-fold higher than that of MMCK from several species. In C. aceratus MCK-1 and MCK-2 lacked one of the lysine residues, Lys 8 and Lys24, respectively,

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comprised the weak interacting lysine pair (Hornemann, Stolz, & Wallimann, 2000). It was important that sequence differences of CK isoforms in icefish myotomal muscle microcompartmentalization for different enzymes’ function (Winnard, Cashon, Sidell, & Vayda, 2003). Comparing the primary sequence differences of homeothermal (rabbit, RM-CK) and poikilothermal (carp, M1-CK) M-CK (Fig. 6A), there were four peptides nearby active site of M-CK, residues 267, 268, 270, and 309 (Fig. 6B). Activity of RM-CK G268N, that changed residue 268 from glycine to asparagine, was raised at low temperature and high pH (10°C, pH 8.0). A dipole–dipole interaction between Ser 285 and Cys 283 might help active site shrink and guarantee substrate interaction (Wu, Li, et al., 2011) (Fig. 7). In order to realize the possible characteristic of residue 268, different replacements of certain residue were generated. It was concluded that a hydrophilic residue, such as asparagine, lysine, or threonine, could enhance RM-CK activity at low temperature and high pH. The water preservation molecules, such as sucrose, could enhance enzyme, with hydrophobic surface residue, activity at low temperature (Wu, Li, Wu, & Hui, 2014). At the same time, some strains of carp M1-CK from higher latitudes contained hydrophilic residues rather than hydrophobic ones on the surface of M1-CK from lower latitudes. Besides, surface amino acids with polar side-chains are also found in teleost LDHs’ family (Fields, Dong, Meng, & Somero, 2015). It can be concluded that M-CK surface hydrophobicity is an important criterion for enzyme activity at low temperature. To sum up, CK provides energy in muscle cells. A transgenic fish with exogenous M-CK is more cold tolerance than wild type fish (Fig. 8). Since there is highly homology primary sequence and secondary structure, M-CK tertiary structure majorly controls its activity at low temperature. Because most of effective differences of M-CKs locate on the surface of molecules, interaction between protein and water molecules is an important point to analysis enzyme activity at low temperature. It is reasonable to study the solvent and solutes influence on enzyme structure and stability. A transgenic M3-CK zebrafish has been verified as a cold-adapted individual (Wang et al., 2014; Wu, Lin, et al., 2011). It is also applied in the carp M1-CK transgenic zebrafish. At the same time, a specific gene polymorphism in M1-CK has been found in cold-adapted carp from Amur River. CK activity at low temperature may majorly adjust cold tolerance ability of carp and other fishes.

Fig. 6 (A) Peptides sequences alignment of CKs. Carp (Cyprinus carpio) M1-CK, M2-CK, and M3-CK were compared with Amur River carp M1-CK (Cyprinus carpio haematopterus) and icefish (Cyprinus carpio haematopterus) MCK1 and MCK2. Most of sequence variation located at residues 1–50. (B) Molecular modeling of carp M1-CK. Cys 283 is the active site and residues 267, 268, 270, and 309 may be the important residues of M form CK to adapt at low temperature.

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A

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Fig. 7 (A) Rabbit M form CK fine structure of active site which Ser 285 and Cys 283 formed a dipole interaction to enlarge the active site which is not formed at Carp M1-CK (B).

Fig. 8 A M3-CK transgenic zebrafish has higher swimming ability at low temperature. Adapted from Wu, C. L., Lin, T. H., Chang, T. L., Sun, H. W., Hui, C. F., & Wu, J. L. (2011). Zebrafish HSC70 promoter to express carp muscle-specific creatine kinase for acclimation under cold condition. Transgenic Research, 20(6), 1217–1226.

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4. FATTY ACIDS DESATURASE Thermal compensation was an important physiological response in poikilothermal species, such as fish, that must accommodate to seasonal or diurnal fluctuations in temperature (Hazel, 1995). Environmental changes in temperature-induced phenotypic plasticity of fish generally required a period of adjustment to modify its biochemical composition, i.e., lipid composition of cell membrane. Lipid composition was diverse in different fish to fashion membranes with physical properties appropriate to the prevailing ambient temperature. The fluctuation of membrane lipid composition was on of the most ubiquitous and few continuously graded cellular responses to temperature change. It was different in fatty acids compositions of fish after adaptation to environmental temperature change (Farkas et al., 1984; Hazel, 1979, 1984; Wodtke, 1978, 1981). On the other hand, “homeoviscous adaptation” hypothesis was a widely accepted model to explain the temperature-dependant physiological adaptation of cell membrane (Macdonald & Cossins, 1985). Since membrane fluidity was directly related to ions and oxygen transportation, changes occurred in the mitochondrial membrane including increase of n-3 and n-6 unsaturated fatty acids (UFAs) contents (Wodtke, 1978) and decline of α-linolenic acid level (Leaver et al., 2006). Previous studies indicated that changes in fatty acids composition in cellular or mitochondrial membrane played a major role in determining cold tolerance in fish (Leaver et al., 2006). Desaturase is the key enzyme for saturated fatty acids (SFAs) desaturation. Stearoyl-CoA desaturase (SCD, EC 1.14.99.5) is the most important enzyme in desaturase enzyme family. It catalyzes stearic acid to oleic acid with transforms NADH to NAD+ (Fig. 9). In physicochemical analysis of different double bond position of fatty acids, double bond in middle site of molecules affected lipid transition temperature obviously

Fig. 9 SCD catalyzes stearoyl-CoA to oleyl-CoA companied by NADH to NAD+, and the reverse reaction.

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(Barton & Gunstone, 1975). The activity of SCD significantly increases and was correlated to membrane fluidity during thermal compensation (Trueman, Tiku, Caddick, & Cossins, 2000). The SCD activity of hepatic microsomes in milkfish significantly increased 2.3-fold at 15°C in comparison to 25°C (Hsieh & Kuo, 2005). Since SCD was found tightly bound to microsomal membranes and endoplasmic reticulum (Man, Miyazaki, Chu, & Ntambi, 2006; Murata & Wada, 1995; Stukey, Mcdonough, & Martin, 1990), its histidine-rich region was thought to be necessary for the protein to bind to iron atoms that participate in the reaction that the enzyme catalyzes (Marquardt, Stohr, White, & Weber, 2000). To compare the different residues of SCD between milkfish, common carp, and grass carp, there was some variation at N-terminal (Hsieh, Liao, & Kuo, 2001). In whole peptide chain there were 12 residues changed to hydrophobic ones, from milkfish to common carp, and 8 residues changed to hydrophilic ones. That might due to the hydrophobic environment of SCD activation. Both temperature and oil-enriched diet could affect the SCD activity in tilapia (Hsieh, Hu, Hsu, & Hsieh, 2007). This study suggested that exposure to low temperatures led to a reduction in the proportion of SFAs and a corresponding increase of UFAs which depended on SCD activity. Recently, mammalian SCD crystal structure has been dissolved (Wang et al., 2015). Further studies on structure aspect of SCD should proceed to evaluate enzyme reaction mechanism. In common carp, there were two isoforms of SCD, which are SCD1 and SCD2. These two isoforms have 93% identity. Carp SCD1 expresses at higher temperature (30°C) and SCD2 at low temperature (less than 17°C). It seemed that SCD isoform expression at different temperature response for biochemical reaction in speculate environment (Polley et al., 2003). SCD protein contained three histidine boxes, two transmembrane regions, and one N-terminal cytochrome b5 domain containing heme-binding motif (Song et al., 2015). There are not so much studies on the relationship between enzyme activities and structured in different temperatures. In SCD study, gene expression level majorly controlled its metabolism efficiency of fatty acid. The regulation of SCD by polyunsaturated fatty acids (PUFAs) and cholesterol were discussed in 1999 (Ntambi, 1999). SREBP and PUFA-BP directly controlled the expression of SCD in mouse model. However, due to highly variation of fish physiological conditions, further studies are still needed to verify the mechanism in fish.

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SCD induced by coconut oil

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Fig. 10 Coconut oil-induced SCD expression after cold treatment for increasing unsaturated fatty acid content in tilapia. Adapted from Hsieh, S. L., Hu, C. Y., Hsu, Y. T., & Hsieh, T. J. (2007). Influence of dietary lipids on the fatty acid composition and stearoyl-CoA desaturase expression in hybrid tilapia (Oreochromis niloticus  O. aureus) under cold shock. Comparative Biochemistry and Physiology Part B: Biochemistry & Molecular Biology, 147(3), 438–444.

Selection of highly expressed SCD milkfish has been applied in aquaculture (Hsieh & Kuo, 2005). It is also the same observation in carp (Balla & Hermesz, 2009). To select a cold tolerance strain in aquaculture is an efficient strategy in aquaculture application. On the other hand, a cold-induced SCD expression is applied to enrich UFAs in tilapia (Fig. 10) (Hsieh et al., 2007).

5. CONCLUSIONS In order to realize the enzyme function of teleost at low temperature, some strategies have been evaluated. Many studies conduct the regulation of genes expression and enzymological mechanism. LDH and CK are the most attractive targets to study temperature depending mechanism of enzyme reaction. On the other hand, SCD is considered as a gene regulation mechanism model for fish at low temperature, due to its critical role of desaturated fatty acids synthesis. In these studies, it may be concluded that activity of enzymes at low temperature depend on the interaction between protein and microenvironment. Hydrophobicity of protein involved in the structural change of certain enzymes. For a soluble enzyme, more hydrophilic residues around protein surface decrease the thermal stability and increase

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flexibility of enzyme may increase its activity and decrease the Gibb’s free energy of enzyme reaction. Furthermore, secondary metabolites which are directly modified by the cellular microenvironment to get adapted to low temperature. Genes of lipid metabolism are major characters of membrane fluidity and hydrophobicity regulators. To study the mechanism and expression profile of fatty acid synthesis and modification enzyme is an important field to construct microenvironment of fatty acid composition.

ACKNOWLEDGMENT We acknowledge the supporting of Academia Sinica, Taiwan in this article.

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Nishiguchi, Y. (2008). Evolutionary implications of lactate dehydrogenases (LDHs) of hagfishes compared to lampreys: LDH cDNA sequences from Eptatretus burgeri, Paramyxine atami and Eptatretus okinoseanus. Zoological Science, 25(5), 475–479. Ntambi, J. M. (1999). Regulation of stearoyl-CoA desaturase by polyunsaturated fatty acids and cholesterol. Journal of Lipid Research, 40(9), 1549–1558. Petricorena, Z. L., & Somero, G. N. (2007). Biochemical adaptations of notothenioid fishes: Comparisons between cold temperate South American and New Zealand species and Antarctic species. Comparative Biochemistry and Physiology. Part A, Molecular & Integrative Physiology, 147(3), 799–807. Pikaard, C. S. (2006). Cell biology of the Arabidopsis nuclear siRNA pathway for RNAdirected chromatin modification. Cold Spring Harbor Symposia on Quantitative Biology, 71, 473–480. Polley, S. D., Tiku, P. E., Trueman, R. T., Caddick, M. X., Morozov, I. Y., & Cossins, A. R. (2003). Differential expression of cold- and diet-specific genes encoding two carp liver delta 9-acyl-CoA desaturase isoforms. American Journal of Physiology. Regulatory, Integrative and Comparative Physiology, 284(1), R41–R50. Sauer, D. M., & Haider, G. (1977). Enzyme activities in the serum of rainbow trout, Salmo gairdneri Richardson; the effects of water temperature. Journal of Fish Biology, 11(6), 605–612. Somero, G. N. (1973). Thermal modulation of pyruvate metabolism in the fish Gillichthys mirabilis: The role of lactate dehydrogenases. Comparative Biochemistry and Physiology. Part B, Biochemistry & Molecular Biology, 44(1), 205–209. Somero, G. N., & Low, P. S. (1976). Temperature: A “shaping force” in protein evolution. Biochemical Society Symposia, 41, 33–42. Song, Y. F., Luo, Z., Pan, Y. X., Zhang, L. H., Chen, Q. L., & Zheng, J. L. (2015). Three unsaturated fatty acid biosynthesis-related genes in yellow catfish Pelteobagrus fulvidraco: Molecular characterization, tissue expression and transcriptional regulation by leptin. Gene, 563(1), 1–9. Stukey, J. E., Mcdonough, V. M., & Martin, C. E. (1990). The OLE1 gene of Saccharomyces cerevisiae encodes the delta 9 fatty acid desaturase and can be functionally replaced by the rat stearoyl-CoA desaturase gene. Journal of Biological Chemistry, 265(33), 20144–20149. Sun, H. W., Hui, C. F., & Wu, J. L. (1998). Cloning, characterization, and expression in Escherichia coli of three creatine kinase muscle isoenzyme cDNAs from carp (Cyprinus carpio) striated muscle. Journal of Biological Chemistry, 273(50), 33774–33780. Sun, H. W., Liu, C. W., Hui, C. F., & Wu, J. L. (2002). The carp muscle-specific subisoenzymes of creatine kinase form distinct dimers at different temperatures. Biochemical Journal, 368(Pt. 3), 799–808. Trueman, R. J., Tiku, P. E., Caddick, M. X., & Cossins, A. R. (2000). Thermal thresholds of lipid restructuring and delta 9-desaturase expression in the liver of carp (Cyprinus carpio L.). Journal of Experimental Biology, 203(Pt. 3), 641–650. Verde, C., Giordano, C., & Di Prisco, G. (2008). The adaptation of polar fishes to climatic changes: Structure, function and phylogeny of haemoglobin. IUBMB Life, 60(1), 29–40. Wang, H., Klein, M. G., Zou, H., Lane, W., Snell, G., Levin, I., et al. (2015). Crystal structure of human stearoyl-coenzyme A desaturase in complex with substrate. Nature Structural & Molecular Biology, 22(7), 581–585. Wang, Q., Tan, X., Jiao, S., You, F., & Zhang, P. J. (2014). Analyzing cold tolerance mechanism in transgenic zebrafish (Danio rerio). PLoS One, 9(7), e102492. Winnard, P., Cashon, R. E., Sidell, B. D., & Vayda, M. E. (2003). Isolation, characterization and nucleotide sequence of the muscle isoforms of creatine kinase from the Antarctic teleost Chaenocephalus aceratus. Comparative Biochemistry and Physiology. Part B, Biochemistry & Molecular Biology, 134(4), 651–667. Wodtke, E. (1978). Lipid adaptation in liver mitochondrial membranes of carp acclimated to different environmental temperatures: Phospholipid composition, fatty acid pattern and cholesterol content. Biochimica et Biophysica Acta, 529(2), 280–291.

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CHAPTER SEVEN

Usefulness of Alginate Lyases Derived from Marine Organisms for the Preparation of Alginate Oligomers with Various Bioactivities S. Takeshita*,1, T. Oda† *Center for Industry, University and Government Cooperation, Nagasaki University, Nagasaki, Japan † Graduate School of Fisheries Science & Environmental Studies, Nagasaki University, Nagasaki, Japan 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Alginate Lyase 2.1 Mannuronate Lyase (PM Lyase) 2.2 Guluronate Lyase (PG Lyase) 2.3 Bifunctional Alginate Lyase 3. Bioactivities of Enzymatically Digested Alginate Oligomers 3.1 Activation of Immune Systems 3.2 Growth-Promoting Effects on Plant Cells References

138 140 142 148 151 153 153 155 157

Abstract Alginate-degrading enzyme, alginate lyase, catalyzes the cleavage of glycosidic 1–4 O-linkages between uronic acid residues of alginate by a β-elimination reaction leaving a 4-deoxy-L-erythro-hex-4-ene pyranosyluronate as nonreducing terminal end. The enzymes from a wide variety of sources such as marine molluscs, seaweeds, and marine bacteria have been discovered and studied not only from a point of view of enzymological interest of enzyme itself but also for elucidation of fine chemical structure of alginate, structure–activity relationship of alginate, and biological activities and physicochemical features of the enzymatic digestion products. Based on the substrate specificities, alginate lyases are classified into three groups: poly(β-D-mannuronate) lyase, poly(α-L-guluronate) lyase, and bifunctional alginate lyase, which are specific to mannuronate, guluronate, and both uronic acid residues, respectively. We have studied enzymological aspects of these three types of alginate lyases, and bioactivities of enzymatically digested alginate oligomers. In this chapter, we described the purification and

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characterization of three types of alginate lyases from different marine origins and overviewed the bioactivities of alginate oligomers.

1. INTRODUCTION Alginate is a linear acidic polysaccharide comprising β-D-mannuronate (M) and its C-5 epimer α-L-guluronate (G). In alginate polymers, the residues are arranged in a block structure of a homopolymer (polymannuronate (PM) or polyguluronate (PG)) or heteropolymer (a mixed sequence of these residues). The block structures are expressed as M-block, G-block, and MG-block (Haug, Larsen, & Smidsrod, 1967; Fig. 1A). The arrangement of these block structures and the molecular size of alginate polymers, which differ depending on the sources, influence the physicochemical characterization such as gel-forming property and viscosity. The G-block binds divalent cations, especially calcium ions strongly, and can contribute to form strong but reversible alginate gels with so-called egg-box structure, whereas M-block or MG-block is involved in weak and relatively elastic gel formation (Murata et al., 1993; Fig. 1B). Alginate is mainly found in brown seaweeds as an intracellular material and comprises about 30% of dry weight of these seaweeds (Gacesa, 1992). Certain bacteria also produce alginate as an extracellular polysaccharide to protect them from detrimental environmental factors. Commercially available alginates are generally prepared from extracts of large brown algae such as Laminaria hyperborea, Macrocystis pyrifera, Laminaria japonica, and other species at industrial levels (Zhu & Yin, 2015). Alginates are widely used in the commercial food and pharmaceutical industries such as stabilizer, viscosifier, gelling agent, film, and therapeutic agents (Gacesa, 1988). Alginate oligosaccharides (oligomers) with relatively low molecular weight are depolymerization products of alginate by either enzymatic degradation or physicochemical methods including acid hydrolysis. It has been demonstrated that some of the alginate oligomers show growth-promoting effects on certain higher plant species and therapeutic effects in mammalian systems such as anticoagulants and antitumor activities (An et al., 2009; Hu, Jiang, Hwang, Liu, & Guan, 2004; Iwamoto et al., 2005). These findings suggest that alginate oligomers, especially the enzymatic-digested alginate oligomers, are capable to exhibit significant valuable biological activities in both plant and animal systems. Our previous study demonstrated that alginate lyase-digested alginate oligomers promoted the growth of freshwater green microalga Chlamydomonas

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Fig. 1 (A) Chemical structures of mannuronate (M)-block, guluronate (G)-block, and MG-block. (B) Schematic structure of alginate gels with so-called egg-box structure. The circles represent calcium ions, and the lines are alginate polymers.

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reinhardtii, while acid-hydrolyzed alginate oligomers had no such growthpromoting activity (Yamasaki et al., 2012). In addition, a significant increase in intracellular fatty acid levels of C. reinhardtii cultured with enzymatically digested alginate oligomers was observed. These findings suggest that alginate lyase-digested alginate oligomers can be used as a promoting agent for efficient biomass production by reducing culture periods and changing fatty acid levels, which might be potentially useful as a source of biofuel (Ueno & Oda, 2014). Furthermore, it has been reported that enzymatically depolymerized alginate oligomers stimulated mouse macrophage cell line RAW264.7 cells to induce cytokine secretion in structure- and concentration-dependent manners, while the activities of acid-hydrolyzed alginate oligomers with same molecular sizes were quite low as compared to the enzyme-degraded oligomers (Iwamoto et al., 2005). These results suggest that preparation procedure of alginate oligomers is important to obtain bioactive oligomers rather than simply molecular size and composition. Probably enzyme-digested alginate oligomers with nonreducing double bond end structure have superior biological activities than those of acid-hydrolyzed alginate oligomers. In this chapter, we would like to describe the purification and characterization of some alginate lyases derived from marine organisms and overview the bioactivities of enzymatically digested alginate oligomers.

2. ALGINATE LYASE Alginate lyases are a group of enzymes which can recognize alginate as a substrate. Aqueous solution of alginate polymer generally has high viscosity and even forms gels in the presence of calcium ion. Hence, the enzymatic activity of alginate lyase can be easily detected by decrease in the viscosity of alginate solution. Therefore, the activity of alginate lyase can be measured with Ostwald’s viscometer (Fig. 2). Plaque formation assay using alginate containing alga plate has also been developed for screening of certain bacteria that produce alginate-degrading enzyme (Takeshita, Oda, & Muramatsu, 1991; Fig. 3). Enzymological studies revealed that alginate lyase catalyzes a β-elimination reaction that splits the 1,4-linkages leading to formation of an uronic acid residue with unsaturated double bond at terminal sugar. Since the double bond has absorbance at 235 nm, alginate lyase with such activity can be measured by the increase in absorbance at 235 nm with the progression of enzymatic reaction (Fig. 4). Marine molluscs such as abalone and turban shell which generally eat marine seaweed containing

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Fig. 2 Time-course of the change in viscosity of alginate solution coursed with endolyticaly reactive alginate lyase.

Fig. 3 Plaque-forming assay for screening of alginate lyase-producing bacteria. Cultured supernatant of an isolated bacterial strain was added to a cup putted on the alga plate containing alginate (A), PG (B), or PM (C) as a substrate, and then the plates were incubated under suitable conditions. To form precipitate of each alginate substrate in the alga, 70% ethanol containing 1% CaCl2 (A and B) or 2 N H2SO4 (C) was added. Due to specific enzymatic degradation, clear plaque was observed around the cup. Plaque formation indicates specific activity of alginate lyase (A), PG lyase (B), and PM lyase (C).

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Fig. 4 Spectrophotometric assay for alginate lyase. PG solution (0.1%) in glycine buffer (pH 9.0) was incubated with PG lyase at 30°C for the indicated periods of time, and absorbance at 235 nm was measured continuously.

alginate as a principal food are easily expected to have alginate digestion enzymes. In fact, pioneering studies on the alginate lyases isolated from digestive cecum of turban shell have been conducted (Muramatsu, Hirose, & Katayose, 1977). In addition to the enzymes isolated from marine molluscs, enzymes with alginate digestion activity have been isolated from various seaweeds and marine bacteria. Enzyme specific for linkages between mannuronic acid residues in alginate polymers, poly(β-D-mannuronate) lyase (PM lyase), has been well studied and [EC 4.2.2.3] has been assigned to this kind of enzyme. On the other hand, [EC 4.2.2.11] has been assigned to enzyme specific for guluronic acid linkages in alginate, poly(α-L-guluronate) lyase (PG lyase). Although EC number has not been assigned yet, bifunctional alginate lyases which can recognize both mannuronic acid and guluronic acid of alginate polymers have been isolated from several marine bacteria. It is considered that such bifunctional alginate lyases are more useful than PM lyase and PG lyase when it comes to efficient preparation of alginate oligomers from various alginate polymers with different M/G ratios (Fig. 5). We describe the details of each of three alginate lyases in terms of their purification and characterization in the following sections.

2.1 Mannuronate Lyase (PM Lyase) 2.1.1 Purification Alginate lyase activity is often detected in digestive fluid of herbivorous marine invertebrates such as abalone, sea hare, and turban shell. Since

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Fig. 5 Substrate-specific actions of PM lyase (A), PG lyase (B), and bifunctional lyase (C). Note that the half of degradation products have nonreducing double bond end structure.

alginate is one of the most abundant carbohydrates in seaweed especially brown algae, it is considered that alginate lyase plays an important role for the assimilation of alginate in these animals. To date many alginate lyases have been identified and characterized. Interestingly, many of the marine molluscan enzymes are classified as PM lyase (Elyakova & Favorov, 1974; Favorov, Vozhova, Denisenko, & Elyakova, 1979; Nakada & Sweeny, 1967; Nisizawa, Fujibayashi, & Kashiwabara, 1968; Rahman, Inoue, Tanaka, & Ojima, 2010; Suzuki, Suzuki, Inoue, & Ojima, 2006). In digestive fluid of gastropods, multiple isozymes of PM lyases with different molecular weights, physicochemical properties, and substrate specificities have been discovered. For instance, recent study reported that two alginate

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lyase isozymes were isolated from sea hare Aplysia kurodai, and these isozymes were characterized as endolytic PM lyase (EC 4.2.2.3) (Rahman et al., 2010). A. kurodai is generally known to feed various different brown seaweeds which may contain alginates with different structural characteristics. Hence, the presence of multiple alginate lyases may be a strategy of this animal for the efficient digestion of alginates with different structures from various seaweeds. In addition to marine invertebrates, some marine bacteria, which are capable of producing PM lyases, have been isolated (Romeo & Preston, 1986; Tseng, Yamaguchi, Nishimura, & Kitamikado, 1992). One of such PM lyase-producing bacterium has been isolated from decayed brown alga (L. japonica), and the bacterium was classified as Vibrio sp. (Wang et al., 2006). Purification and characterization of the enzyme produced by this bacterial strain indicated that the enzyme is highly specific to polymannuronate (PM lyase), and the molecular weight and optimal pH and temperature were different from those of marine invertebrates. Especially the estimated molecular weight (62.5 kDa) was significantly higher than others. Based on the molecular masses, alginate lyases seem to be grouped into three types: small (25–30 kDa), medium (around 40 kDa), and large size (>60 kDa) (Osawa, Matsubara, Muramatsu, Kimura, & Kakuta, 2005). Thus, the large molecular size of the bacterial alginate lyase is a characteristic feature, although the biochemical significance of the large molecular size is still unclear. Purification procedures of enzymes generally depend on the sources and biochemical features of the enzymes, and it is established by repeated try and errors. As an example, we show a purification procedure of PM lyase from midgut gland of turban shell Turbo cornutus. First, midgut gland was removed from live turban shell, and homogenized with phosphate buffer, pH 6.0. After ammonium sulfate precipitation from 50% to 100% saturation of the crude extract, the fraction was subjected to column chromatography using SP-Sephadex C-50, subsequent Sephadex G-100, and finally two times of Bio-Gel P-150 (Muramatsu et al., 1977). During the chromatography, the enzyme activity was detected by increase in absorbance at 235 nm, and separated two peaks were found on SP-Sephadex C-50 column chromatography in the presence of ethylenediaminetetraacetic acid disodium salt. These results suggest that chromatographically distinguishable two fractions existed in the extract of midgut gland of T. cornutus. Since the two fractions were finally obtained as highly purified forms, these were designated SP1 and SP2 as isozymes of PM lyase (Muramatsu & Egawa, 1980).

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2.1.2 Characterization The molecular weight of the alginate lyase SP1 was estimated to be approximately 25,000 by ultracentrifugation analysis. SP1 was most active at a pH range of about 8.8–9.2 and most stable at pH 5–6. Phosphate showed strong stabilizing and enhancing effects on the enzyme activity (Muramatsu et al., 1977). The initial viscosity of alginate solution (0.2%) in Tris–HCl buffer at pH 8.0 decreased sharply by 70% within 5 min immediate after the addition of SP1, suggesting that SP1 acts as endolytic reaction enzyme (Muramatsu & Katayose, 1979). On the other hand, the profiles of the optimal pH, pH stability, thermal stability, and molecular size of SP2 were almost the same as those of SP1. SP2 also caused a rapid decrease in the viscosity of alginate solution with similar kinetics of those of SP1. The isoelectric points of SP1 and SP2 were 7.5 and 7.7, respectively (Muramatsu & Egawa, 1980). The amino acid compositions of SP1 and SP2 were slightly different from each other (Table 1). The total number of amino acid residues in SP1 and SP2 is 252 and 255, respectively. Both SP1 and SP2 are consisted of single polypeptide, and contain only one cysteine residue located inside the molecule, and contain two intramolecular disulfide bonds per polypeptide. Chemical modification analysis suggests that one particular tryptophan residue out of nine residues in SP1 and SP2 seems to be essential for the alginate lyase activity. Further analysis suggests that 1 lysine residue out of 14 residues in SP1 and SP2 probably serves as the functional group with other essential amino acid residue(s) at the active site of the enzyme molecule (Muramatsu & Egawa, 1982). Circular dichroic spectra and optical rotatory dispersions suggest that the entire shape of the protein molecule of both isozymes is spherical, and the secondary structure is most likely β-sheet (Muramatsu, Hashimoto, & Takahashi, 1984). Fig. 6A shows the circular dichroic spectrum of SP2. The pattern of the spectrum reflects the presence of typical β-sheet structure. To analyze the substrate recognition mechanism of the enzymes, highly purified mannuronate oligomers with varying degree of polymerization (DP) were prepared. Using the purified oligomers with defined DP, it was found that both SP1 and SP2 recognize mannuronate tetramer or longer oligomers with higher affinity than oligomer with lower DP, and the span of substrate recognition site in the active site of both enzymes was estimated to be five mannunorate residues (Fig. 7A; Muramatsu, Yamada, Date, & Yoshioka, 1993). Detailed structural analysis on SP1 and SP2 was conducted and found that the amino acid sequences of SP1 and SP2 were almost identical to each other expect for two hydrophobic C-terminal amino acid residues of SP2, isoleucine and leucine, which

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Table 1 Amino Acid Compositions of Alginate Lyases from Various Origins Residues per Molecule PM Lyasea Amino Acid

SP1

SP2

PG Lyaseb

PG Lyasec

Bifunctional Lyased

Lys

14

14

13

10

10

His

16

16

8

8

4

Arg

10

10

13

8

6

Asx

36

36

51

49

38

Thr

15

16

26

13

22

Ser

25

25

38

24

25

Glx

11

11

35

26

23

Pro

9

9

11

6

7

Gly

28

28

23

35

20

Ala

7

7

16

11

8

Val

15

15

17

19

20

Met

3

3

2

2

4

lle

7

8

15

13

10

Leu

13

14

23

15

11

Tyr

13

13

16

19

9

Phe

16

16

10

7

13

Trp

9

9

5

11

1

Cys

5

5

4

2

2

a

PM lyase isozymes isolated from Turbo cornutus. PG lyase isolated from Vibrio sp. PG lyase isolated from Pseudomonas sp. d Bifunctional alginate lyase isolated from Pseudoalteromonas sp. b c

were absence in SP1. The molecular weight of SP2 was calculated to be 28,912 from the amino acid sequence. Two intramolecular disulfide bonds were found to be between 106 and 115 and between 145 and 150 amino acid residues, and a free SH group was located at 236 in these enzymes. Interestingly, SP2 showed stronger heat stability than SP1. Hence, it is speculated that extra C-terminal two amino acid residues in SP2 might contribute to keeping the entire conformation of enzyme molecule required for the full

Fig. 6 Circular dichroic spectrum of PM lyase (SP2) from Turbo cornutus (A) and PG lyase from Vibrio sp. (B).

Fig. 7 Schematic models of substrate recognition site of PM lyase (SP1 and SP2) from Turbo cornutus (A) and bifunctional alginate lyase from Pseudoalteromonas sp. No. 272 (B). Arrows indicate supposed cleavage site of recognized alginate substrates with enzymes.

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enzyme activity. Although the biological significance of the presence of SP1 and SP2 in the midgut gland of T. cornutus is still unclear, these findings on structural aspects of SP1 and SP2 may provide tips for the structure–activity relationship of PM lyase (Muramatsu et al., 1996).

2.2 Guluronate Lyase (PG Lyase) 2.2.1 Purification Several marine bacterial strains have been isolated as alginate lyaseproducing bacteria so far, and it seems that many of them are classified into Pseudomonas and Vibrio genus (Wong, Preston, & Schiller, 2000). In addition to major group of bacterial stains with an ability of producing only single kind of alginate lyase, Pseudomonas fluorescens is known to produce multiple enzymes with different enzymological characteristics simultaneously (Li, Jiang, Guan, Wang, & Guo, 2011). Many of the alginate lyases from bacterial origin are known to degrade G-block and MG-block. In fact, PG lyase has been mainly isolated from marine bacteria (Boyen, Bertheau, Barbeyron, & Kloareg, 1990; Brown & Preston, 1991; Kitamikado, Tseng, Yamaguchi, & Nakamura, 1992; Kobayashi, Uchimura, Miyazaki, Nogi, & Horikoshi, 2009; Li, Jiang, et al., 2011; Ostgaard, Knutsen, Dyrset, & Aasen, 1993; Shimokawa et al., 1997). In the case of P. fluorescens which was isolated from decayed brown seaweed in the Yellow Sea of China (Li, Jiang, et al., 2011), it has been reported that the bacterial strain produced three alginate lyases with different substrate specificity (A, B, and C). Enzymes A and B showed the activity for both M- and G-blocks, while enzyme C was highly specific to G-block. As a unique alginate lyaseproducing bacterium, a deep-sea bacterium Agarivorans sp. has been reported to produce high-alkaline alginate lyase which preferably degraded MG- and G-rich fragments in alginate (Kobayashi et al., 2009). More recently, a novel alginate lyase has been obtained from the polysaccharide-degrading bacterium Flammeovirga sp. strain MY04. This enzyme prefers guluronate (G) to mannuronate (M) with endolytic degradation manner, which is a 566 amino acid protein and belongs to a novel branch of the polysaccharide lyase 7 (PL7) superfamily. The smallest substrate for the enzyme is an unsaturated pentasaccharide, and the enzymedigested products are disaccharides to heptasaccharides. Its minimum product is an unsaturated disaccharide. Since alginate lyase cleavages double bond between C-4 and C-5 of adjacent sugar residues at C-4 position, the enzyme degradation products contain unsaturated nonreducing terminus (Fig. 5), which is usually designated as ΔM or ΔG to distinguish from the G and

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M residues without double bond. Interestingly, the disaccharides produced by alginate lyase of Flammeovirga sp. strain MY04 are identified as ΔG units, while the tri- and tetrasaccharides contain higher proportions of ΔG to ΔM ends. Moreover, the larger final products are only ΔM ends (Han et al., 2016). We have screened for PG lyase-producing bacteria from marine environments. For the screening, it is necessary to establish an efficient detecting method for PG lyase-producing bacteria. We could finally develop a useful screening method (Takeshita et al., 1991). Briefly, 0.1% guluronate polymer was added to usual agar medium for screening of PG lyase-producing bacteria. After cultivation of target bacteria for a while on the alga plates, the plates were immersed in 1% calcium chloride together with 70% ethanol to precipitate undigested PG in the medium. As a result, clear plaque was formed around the colonies of lyase-positive bacteria due to the degradation of PG which does not precipitate as mentioned earlier (Fig. 3). By this screening method, we could discover several PG lyase-producing bacterial strains. A marine bacterium, Vibrio sp. isolated from intestinal contents of a red sea bream (Pagrus major) captured in Tachibana Bay, Nagasaki, Japan, was found as a highly potent producer of PG lyase (Takeshita, Morii, Yoshikoshi, Oda, & Muramatsu, 1995). The isolated bacterial strain grew well in a medium containing 0.5% polypeptone, 0.1% yeast extract, 0.01% KH2PO4, 3.0% NaCl, 0.07% KCl, 0.26% MgSO4, 0.5% MgCl2, and 0.2% sodium alginate (pH 7.5). To characterize the enzyme produced by this bacterial strain, the cultured supernatant was fractionated with ammonium sulfate precipitation from 50% to 100% saturation, and then the precipitated fraction was subjected to column chromatography of DEAE-Cellulofine twice. Subsequently the active fraction was applied to Cellulofine GCL1000m and Butyl-Cellulofine column chromatography continuously. By these chromatographical procedures, highly purified PG lyase was eventually obtained (Takeshita, Sato, Igarashi, & Muramatsu, 1993). Another example of PG lyase-producing bacterial strain was classified into Pseudomonas sp. which was named as Pseudomonas sp. strain F6. This strain grew well in medium containing 0.2% sodium alginate, 0.1% NaNO3, 0.01% KH2PO4, 0.001% FeCl3, and 0.1% Tris (hydroxymethyl) aminomethane in seawater (pH 7.5). From the cultured supernatant of Pseudomonas sp. strain F6, purified enzyme was obtained by similar procedure as described earlier. Namely, ammonium sulfate precipitation and subsequent column chromatography using DEAE-Cellulofine, Sephadex G-100, and Butyl-Cellulofine (Miyazaki, Obata, Iwamoto, Oda, &

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Muramatsu, 2001). Based on these two examples of purification of PG lyase from bacterial origin, it seems likely that purification of PG lyase from culture supernatant of lyase-positive bacteria can be conducted by usual chromatographical procedure without complicated procedure, and these studies may provide useful information regarding the purification procedures for PG lyase from newly isolated bacterial strains. 2.2.2 Characterization The molecular weight of PG lyase from Vibrio sp. was estimated to be 42,000 by SDS-PAGE and of 40,000 by gel-filtration chromatography. The optimal pH and stable pH range for the enzyme activity were 8.5 and 6–10, respectively. As an interesting biochemical feature, it was found that this enzyme was relatively heat stable, and the activity remained without a noticeable decline up to 70°C. Although gradual decrease in the activity was observed beyond this temperature, even at 100°C, the enzyme still kept 45% of the original activity. The maximal activity was attained at 45°C. Moreover, this enzyme showed significant resistant property against the treatment with common protein denaturants. After treatment with 3% SDS, 6 M guanidine hydrochloride (GHCl), or 3 M urea at 25°C for 30 min, the remained enzyme activities were 100%, 70%, or 75% of original native enzyme activity, respectively (Takeshita et al., 1993). To gain a clue for the resistant mechanism of the enzyme against these denaturant agents, the far- or near-ultraviolet circular dichroic spectra of the enzyme in the presence of each denaturant were measured. The results indicate that the secondary structure of the enzyme is hardly affected by treatment with 6 M GHCl but is influenced by 3 M urea and 3% SDS. On the other hand, the near-ultraviolet circular dichroic spectra were attenuated by treatment with 6 M GHCl and 3 M urea, suggesting that the adjacent environment of tryptophan residues in the enzyme molecule may be changed by 6 M GHCl and 3 M urea. After removal of the denaturants by dialysis, far- or near-ultraviolet circular dichroic spectra of the enzyme were restored to almost original spectra without denaturants, suggesting the restoration of native protein conformation. Probably PG lyase is more susceptible to ureainduced denaturation than to GHCl. Since the enzyme activity and conformation were recovered rapidly to original levels under the gentle conditions in the absence of denaturant, it seems that the denaturation may be partially limited to the secondary and/or tertiary structure of the enzyme molecule. This may be due to the rigid core domain of the enzyme rich in β-sheet structure, which is known to be resistant to denaturant-induced

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conformational changes. Fig. 6B shows the circular dichroic spectrum of PG lyase from Vibrio sp. The pattern is similar to that of SP2. These circular dichroic spectra suggest that both enzymes have β-sheet structure as a common structural feature even though the origin of the enzymes is different. The amino acid composition of PG lyase was determined (Table 1). The total number of amino acid residues is 326. The protein has relatively high number of acidic amino acid and serine residues (Takeshita, Oda, & Muramatsu, 1995). The molecular weight of PG lyase from Pseudomonas sp. strain F6 was estimated to be 36,000 by SDS–PAGE. The optimal pH and stable pH range of this enzyme were pH 7.5 and between pH 6.5 and 8.5, respectively. The thermal stability of this enzyme was slightly less than the PG lyase from Vibrio sp. as described earlier. This enzyme also showed relatively resistance against common protein denaturants, although the extent of the resistance against denaturants was less than the PG lyase from Vibrio sp. After the treatment of the enzyme with 1 M GHCl or 4 M urea, approximately half of the original activity still remained, and denaturation with SDS was moderate. Similar to other PM lyase from mollusc and PG lyase from marine bacterium described earlier, this enzyme also has β-structure. Hence, it seems likely that β-structure is a common secondary structure of alginate lyases even from different origins. It has been known that many of alginate lyases reported previously require divalent cations for the maximum activity (Gacesa, 1992). Consistent with the finding, enzyme activity of PG lyase from Pseudomonas sp. strain F6 decreased by 30% of the native enzyme activity in the presence of EGTA, a calcium-specific chelator, and the activity restored effectively by the addition 0.1 M CaCl2, indicating that the enzyme from Pseudomonas sp. also requires a divalent cation, at least calcium ion for the full enzyme activity. The amino acid compositions of this enzyme were determined (Table 1). The total number of amino acid residues is 278. This protein also has a relatively high number of acidic amino acid residues and glycine (Miyazaki et al., 2001).

2.3 Bifunctional Alginate Lyase Conformation of mannuronate and guluronate is quite different each other. From the point of view of classical lock-and-key model for enzyme, it was considered that alginate lyases should be categorized into two groups based on the substrate specificities, PM and PG lyase, and the classification was widely accepted. However, some of the alginate lyases frequently show

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activities toward both PM and PG, although the activities are different depending on the substrates. Some crude enzyme extract shows relatively wide substrate specificities. At first, it is considered that this may be due to the presence of multiple alginate lyases with different substrate specificity in the extract. On the contrary, however, some enzymes have been discovered, which even at highly purified form clearly showed activities against both M-block and G-block (Dou et al., 2013; Li, Dong, et al., 2011; Sawabe, Ohtsuka, & Ezura, 1997; Wang, Guo, Yu, & Han, 2013; Zhu, Chen, Yin, Du, & Ning, 2016; Zhu, Wu, et al., 2016). It is now believed the presence of alginate lyase which can recognize both PM and PG, and degrade them. Our previous studies also showed that a marine bacterium, Pseudoalteromonas sp. strain No. 272 isolated from sea mud in Omura Bay, Nagasaki, Japan, produced an alginate lyase that acts on both M-block and G-block (Iwamoto et al., 2001). 2.3.1 Purification An alginate lyase-positive marine bacterial strain was obtained from mud in Omura Bay, Nagasaki, Japan. The marine bacterium was identified as Pseudoalteromonas sp. strain No. 272 and was cultured in a seawater-based medium containing 0.2% sodium alginate, 0.7% K2HPO4, 0.2% KH2PO4, 0.1% (NH4)2SO4, and 0.05% sodium citrate (pH 8.0). Culture conditions of this bacterial strain were slightly different from those of PG lyase-producing marine bacteria described earlier, but it seems likely that sodium alginate is an essential ingredient for the alginate lyase-producing bacteria. For the purification of the enzyme, the cultured supernatant was saturated with ammonium sulfate to 80% first, and the fraction was applied to column chromatography using DEAE-Cellulofine A500 and Sephadex G-100 (Iwamoto et al., 2001). The established chromatographical purification procedure for this enzyme is also basically the combination of ion-exchange and gelfiltration chromatographic methods. 2.3.2 Characterization The isoelectric point of bifunctional alginate lyase from Pseudoalteromonas sp. strain No. 272 was 3.8. Hence, the enzyme is relatively acidic protein. As a conformational feature of the enzyme, far-ultraviolet circular dichroic spectrum of this enzyme suggested the presence of β-sheet structure. The optimal pH and stable pH range of the enzyme were 7.5–8.0 and 5–11, respectively. In terms of heat stability, the enzyme was more stable in

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phosphate buffer (pH 7.0) than in Tris–HCl buffer (pH 7.0). Although the exact reason for this is still unclear, Tris might act as detrimental factor for this enzyme. Since the enzyme caused immediate decrease in the viscosity of sodium alginate solution, this enzyme is considered to have an endo-type action manner. Analysis using purified alginate oligomers with defined DP revealed that the enzyme did not recognize trimeric guluronate and mannuronate, but efficiently acted on oligomers longer than tetramers. The kinetic analysis suggests that the enzyme from Pseudoalteromonas sp. strain No. 272 recognizes six uronic acid residues (both guluronate and mannuronate oligomers) (Fig. 7B; Iwamoto et al., 2001). The span of substrate recognition site of this enzyme is one reside longer than that of PM lyase from T. cornutus (SP1 and SP2) as seen in Fig. 7A. Probably the most efficiently recognized uronic acid oligomer size differs depending on enzymes. The enzyme consisted of 233 amino acid residues, and the molecular mass was calculated to be 25,549. Chemical modification analysis suggested that lysine, tyrosine, and tryptophan residues might be involved in the enzyme activity in different ways (Iwamoto, Iriyama, Osatomi, Oda, & Muramatsu, 2002). Moreover, a fluorescence-quenching study suggested that the sole tryptophan residue is located in a buried region of the enzyme molecule (Iwamoto, Hidaka, Oda, & Muramatsu, 2003).

3. BIOACTIVITIES OF ENZYMATICALLY DIGESTED ALGINATE OLIGOMERS 3.1 Activation of Immune Systems As an immune-stimulating activity of enzymatically digested alginate oligomers, the activities to induce cytokine secretion from mouse macrophage cell line RAW264.7 cells are described in this section (Iwamoto et al., 2005). Analysis using alginate oligomers with different DPs, which were prepared by enzymatic digestion of PG and PM, showed that the activity of the oligomers to induce TNF-α secretion from RAW264.7 cells differed depending on the oligomer structures. Among the oligomers (M3
M8 and G3
G8) tested, G8 and M7 showed the highest activity (Iwamoto et al., 2005). On the other hand, saturated alginate oligomers (M3
M8 and G3
G8) prepared by acid hydrolysis showed significantly lower activities than those of unsaturated oligomers, suggesting that the unsaturated terminal structure with double bond in the enzymatic-digested alginate oligomers is involved in the bioactivity of alginate oligomers. RAW264.7 cells may recognize unsaturated terminal structure of alginate oligomer. Bio-Plex assay, which

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is a bead assay allowing measurement of multiple cytokines in a single sample simultaneously, demonstrated that G8 and M7 induced IL-1α, IL-1β, and IL-6 production from RAW264.7 cells in addition to TNF-α (Yamamoto, Kurachi, Yamaguchi, & Oda, 2007). Among the oligomers tested, G3 and M3 showed relatively higher cytokine-inducing activities than other oligomers. Based on these findings, it is possible to speculate that trimer such as G3 and M3 is the most suitable molecular size for the specific receptors on RAW264.7 cells, which may be linked with intracellular signaling pathway leading to cytokine secretion. To our knowledge, this is a first study showing that highly purified alginate oligomers can exhibit cytokine-inducing activity depending on the structure. Alginate oligomers, especially enzyme-digested oligomers, are useful not only as immunepotentiating agents but also as a tool for analyzing pattern recognition mechanism of macrophages. Further study of structure–activity relationship of alginate oligomers prepared with appropriate alginate lyases may provide insight into specific receptor-mediated recognition mechanism as well as intracellular signal transduction system leading to the cytokine secretion in macrophages. Comparative study using various alginate polymers with different molecular sizes showed that the activities of original alginate polymers to induce TNF-α secretion from RAW264.7 cells significantly increased after alginate lyase digestion (Kurachi et al., 2005). Furthermore, intraperitoneal (i.p.) injection of alginate lyase-digested alginate oligomer mixture (AOM) into mice resulted in significant increase in several cytokines levels in mice serum (Yamamoto et al., 2007), and the increase in G-CSF (granulocyte colonystimulating factor) level was the most significant. Detailed analysis on G-CSF level in the serum showed that the highest level of G-CSF was detected at 2 h after injection of AOM at 700 mg/kg body weight, and nearly similar high level of G-CSF was maintained until 6 h, and then gradually decreased to basal level thereafter. On the other hand, i.p. injection of original alginate polymer (700 mg/kg) did not induce the increase in G-CSF level. Hence, it is considered that alginate oligomers acquired potent cytokine-inducing activity that was not observed in the original polymer. Exact reason for this is still unclear, but high viscosity and calcium ion-induced gel-forming property of alginate polymer in physiological condition may suppress the approach of alginate polymer to immune system involved in cytokine production. Such physicochemical properties of alginate polymer may be changed by enzymatic degradation, which may lead to the improved pharmacokinetics and became capable of inducing cytokine in in vivo

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system. In general, bioactivities of alginate polymers differ depending on molecular size, M/G ratio, and entire molecular conformation. Thus, it is important to choose appropriate alginate polymers with suitable physicochemical and biological properties depending on the purpose. Nevertheless, extensive variety of structural features of alginate polymers derived from variety of M/G ratios and sequence of M and G residues make the exact characterization of alginate polymers very difficult. As mentioned earlier, bioactivities of enzyme-digested alginate oligomers are much improved as compared to the original polymers. Therefore, alginate lyase digestion of alginate polymers may be a promising strategy to increase the bioactivities of alginate polymers. As an example for the improved bioactivity of alginate oligomers in in vivo mammalian model, it has been reported that i.p. administration of alginate oligomers with average DP of 4.4 resulted in IL-12 production concomitant with the suppression of Th2 development and IgE production, which may lead to antiallergic effect (Yoshida, Hirano, Wada, Takahashi, & Hattori, 2004).

3.2 Growth-Promoting Effects on Plant Cells In addition to mammalian systems as mentioned earlier, alginate oligomers also influence some plant systems. It has been reported that bacterial alginate lyase-digested alginate oligomers promoted shoot elongation of komatsuna (Brassica rapa var. pervidis) seeds (Yonemoto et al., 1993) and the elongation of barley roots (Tomoda, Umemura, & Adachi, 1994). In addition to higher plants, recent studies have demonstrated that enzymatically digested alginate oligomers showed growth-promoting effects on microalgae. In aquaculture industry, Nannochloropsis oculata classified into Eustigmatophyceae classis is a useful marine unicellular microalga, which is widely used as an important feeding food for bivalves, larvae of crustacean, and some fish species. Yokose et al. reported that the growth of N. oculata was promoted by bacterial alginate lyase-digested AOM in a concentration-dependent manner (Yokose et al., 2009). In the presence of 1 mg/mL of AOM, slight growth promotion of N. oculata was observed, and the maximum effect was induced at 20 mg/ mL. Cell number of N. oculata cultured with AOM (20 mg/mL) became almost five times higher than that of control without AOM after reached to stationary growth phase. Since the growth of N. oculata in the presence of 40 mg/mL of AOM was slightly down as compared to the level at 20 mg/mL, there may be an appropriate concentration of AOM for the maximum growth-promoting effect on N. oculata.

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The culture of N. oculata is often suffered from various environment stresses. For example, some heavy metals even at trace level can cause negative effects on microalgae (Satoh, Vudikaria, Kurano, & Miyachi, 2005). In aquaculture industry, copper ion (Cu2+) is often used for disinfection of parasites. However, Cu2+ shows highly toxic effect on microalga. In fact, Cu2+ at 2.5 mg/mL significantly suppressed the growth of N. oculata, and cell division was completely arrested after 24 h incubation. However, concomitant addition of AOM (10 mg/mL) resulted in almost complete disappearance of the toxic effect of Cu2+ on N. oculata. These results suggest that AOM is capable of not only promoting the growth of N. oculata under normal condition but also improving growth environment by reducing the impact of stressors. As far as we know, this is the first study showing that enzymedigested alginate oligomers have growth-promoting effect on valuable marine microalga N. oculata. Considering the safety and environmentally friendly material, alginate oligomers can be introduced to aquaculture farms to improve the culture condition of valuable marine microalgae. The green alga C. reinhardtii occurs globally in many soil and freshwater environments. C. reinhardtii is an important model species for fundamental studies in plant physiology, biochemistry, and molecular biology. C. reinhardtii is also considered as promising alga to produce biofuels. In fact, C. reinhardtii has been used in a number of studies on algal hydrogen production (Yamasaki et al., 2012). Next we would like to show the comparative study on the effects of AOM. In the study, comparison between alginate lyase-digested (ED-AOM) and acid-hydrolyzed alginate oligomer mixture (AH-AOM) was conducted in terms of the effect on the growth of C. reinhardtii. ED-AOM showed significant growth-promoting effect on C. reinhardtii in a concentration-dependent manner, and the maximum effect was observed after 4 days cultivation. Unexpectedly the fatty acid composition of C. reinhardtii was influenced by ED-AOM. Namely, contents of C16:0, C18:2 cis, and C18:3 n-3 increased in treated cells. Timecourse and concentration-dependent analysis indicated that the maximum effect of ED-AOM was attained at 1000 μg/mL on day 4. On the other hand, AH-AOM was obviously less effective as compared to ED-AOM. Recently, microalgae are drawing extensive attention as alternative promising sources such as biofuel and biodiesel (Schenk et al., 2008). Due to the feasibility of mass culture of microalgae, oil components of microalgae may be especially attractive biofuel sources, which can be produced at industrial level. ED-AOM seems to affect fatty acid compositions of C. reinhardtii in addition to the growth promotion. Contents of C16:0, C18:2 cis, and

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C18:3 n-3 were increased in the presence of enzymatically prepared AOM. Furthermore, contents of some other fatty acids such as C14:1, C17:0, and C20:0 were also increased. Although the exact mechanism of the effect of enzymatically prepared AOM on the composition of fatty acids in C. reinhardtii is still unclear, this effect may lead to improvement of the usefulness of green algae.

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Yamamoto, Y., Kurachi, M., Yamaguchi, K., & Oda, T. (2007). Stimulation of multiple cytokines production in mice by alginate oligosaccharides following intraperitoneal administration. Carbohydrate Research, 342, 1133–1137. Yamasaki, Y., Yokose, T., Nishikawa, T., Kim, D., Jiang, Z., Yamaguchi, K., et al. (2012). Effects of alginate oligosaccharide mixtures on growth and fatty acid composition of the green alga Chlamydomonas reinhardtii. Journal of Bioscience and Bioengineering, 113, 112–116. Yokose, T., Nishikawa, T., Yamamoto, T., Yamasaki, Y., Yamaguchi, K., & Oda, T. (2009). Growth-promoting effect of alginate oligosaccharides on a unicellular marine microalga, Nannochloropsis oculata. Bioscience, Biotechnology, and Biochemistry, 73, 450–453. Yonemoto, Y., Tanaka, H., Yamashita, T., Kitabatake, N., Ishida, Y., Kimura, A., et al. (1993). Promotion of germination and shoot elongation of some plants by alginate oligomers prepared with bacterial alginate lyase. Journal of Fermentation and Bioengineering, 75, 68–70. Yoshida, T., Hirano, A., Wada, H., Takahashi, K., & Hattori, M. (2004). Alginic acid oligosaccharide suppresses Th2 development and IgE production by inducing IL-12 production. International Archives of Allergy and Immunology, 133, 239–247. Zhu, B., Chen, M., Yin, H., Du, Y., & Ning, L. (2016). Enzymatic hydrolysis of alginate to produce oligosaccharides by a new purified endo-type alginate lyase. Marine Drugs, 14, 108–118. http://dx.doi.org/10.3390/md14060108. Zhu, Y., Wu, L., Chen, Y., Ni, H., Xiao, A., & Cai, H. (2016). Characterization of an extracellular biofunctional alginate lyase from marine Microbulbifer sp. ALW1 and antioxidant activity of enzymatic hydrolysates. Microbiological Research, 182, 49–58. Zhu, B., & Yin, H. (2015). Alginate lyase: Review of major sources and classification, properties, structure-function analysis and applications. Bioengineered, 6, 125–131.

CHAPTER EIGHT

Marine Microbial Amylases: Properties and Applications J. Suriya*, S. Bharathiraja†, M. Krishnan*, P. Manivasagan{, S.-K. Kim§,1 *School of Environmental Sciences, Bharathidasan University, Tiruchirappalli, Tamil Nadu, India † CAS in Marine Biology, Annamalai University, Porto Novo, Tamil Nadu, India { Marine Bioprocess Research Center, Pukyong National University, Busan, Republic of Korea § Marine Bioprocess Research Center; Specialized Graduate School Science & Technology Convergence, Pukyong National University, Busan, Republic of Korea 1 Corresponding author: e-mail addresses: [email protected]; [email protected]

Contents 1. Introduction 2. Types of Amylase 2.1 α-Amylase 2.2 β-Amylase 2.3 Glucoamylase 3. Production Methods 4. Production of Amylases 4.1 Production of α-Amylase 4.2 Production of β-Amylase 4.3 Production of Glucoamylase 5. Purification of Amylases 6. Characterization of Amylases 7. Molecular Biology of Amylases 8. Determination of Enzyme Activity 8.1 Dinitrosalicylic Acid Method 8.2 Nelson–Somogyi Method 8.3 Determination of Activity Using Iodine 8.4 Dextrinizing Activity 8.5 Indian Pharmacopeia Method 8.6 Reduction in Viscosity of Starch Suspension 9. Industrial Applications Amylases 9.1 Enzymatic Hydrolysis of Starch for Fructose and Glucose Production 9.2 Bakery Industry 9.3 Other Industrial Applications 10. Conclusion Acknowledgment References

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Abstract Amylases are crucial enzymes which hydrolyze internal glycosidic linkages in starch and produce as primary products dextrins and oligosaccharides. Amylases are classified into α-amylase, β-amylase, and glucoamylase based on their three-dimensional structures, reaction mechanisms, and amino acid sequences. Amylases have innumerable applications in clinical, medical, and analytical chemistries as well as in food, detergent, textile, brewing, and distilling industries. Amylases can be produced from plants, animals, and microbial sources. Due to the advantages in microbial production, it meets commercial needs. The pervasive nature, easy production, and wide range of applications make amylase an industrially pivotal enzyme. This chapter will focus on amylases found in marine microorganisms, their potential industrial applications, and how these enzymes can be improved to the required bioprocessing conditions.

1. INTRODUCTION More than 70% of the Earth surface is covered by sea, has unprecedented biological diversity, and comprises 95% of the biosphere (Qasim, 1999). Microbial diversity in marine environment is a promising agent for many novel industrial products (Berdy, 2005; Fenical & Jensen, 2006). The recent researches showed that the culturability of microorganisms in marine sediments (0.25%) and seawater (0.001–0.10%) is lower than soil microorganisms (0.30%) (Amann, Ludwig, & Schleifer, 1995). Henceforth, researchers switched over to marine environments for novel products from microorganisms. Microorganisms inhabitating in marine environment are always under stress conditions for space, nutrient, defence, offence, and salinity. Some marine eco regions are always changing its environmental conditions. Microorganisms living under this environment produce unique characteristic enzymes which are not produced by terrestrial organisms. These enzymes may have more than one independent function and these are considered as multifunctional enzymes. The main advantages of enzymes are the possibility to work at milder conditions, highly specific to substrate as well as eco-friendly (Krishna, 2011). Amylase is an extracellular enzyme which degrades starch by hydrolyzing glycosidic bonds at α-1,4 position (Ajayi & Fagade, 2007) and produces different polysaccharides such as glucose, maltose, and dextrin (Riaz, Haq, & Qadeer, 2003). Amylases have been used since ancient times (Gupta, Gigras, Mohapatra, Goswami, & Chauhan, 2003) for starch

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saccharification, beer production, digestive disorders treatment, and cheese production from milk (Drauz, Gr€ oger, & May, 2012). The enzymatic class of amylase accounts for 25% of world’s market (de Freitas, Escaramboni, Carvalho, de Lima, & de Oliva-Neto, 2014). Cost-effective medium is crucial to meet out the demands of amylase utilizing industries (Balkan & Figen, 2007). Utilization of agro-industrial wastes for the production of amylase enzyme is the cost-effective method and it also solves the problem of pollution (Amin, Bhatti, Zuber, Bhatti, & Asgher, 2015; Singh, Kapoor, & Kumar, 2012). The following sections elaborate on the types of amylases, their production, and their industrial applications.

2. TYPES OF AMYLASE 2.1 α-Amylase α-Amylase (EC 3.2.1.1) hydrolyze the internal α-1,4-glycosidic linkages in starch and produce glucose and maltose. α-Amylase depends on calcium for its catalysis and cannot cleave the terminal glucose residues and α-1,6-linkages. Starch is a polysaccharide that consists of 75–80% of amylopectin and 20–25% of amylase, and it is the natural substrate for the activity of α-amylase. Amylose is a linear chain molecule consisting of glucose units united by α-1,4-glycosidic linkage, whereas amylopectin has branched chains of glucose units. α-1,4-Glycosidic linkage joined the linear glucose units, while α-1,6-glycosidic bonds are present in branched chain which occurs at every 15–45 glucose units. The composition of glucose and maltose obtained after hydrolysis of starch is dependent on the hydrolysis conditions like the temperature and the enzyme origin. pH 7.0 is optimum for the activity α-amylase (Sundarram & Murthy, 2014).

2.2 β-Amylase β-Amylase (EC 3.2.1.2) produces maltose from starch by hydrolyzing the α-1,4-glucan linkages. Seeds of higher plants and sweet potatoes are the primary sources of β-amylase. β-Amylase is responsible for the sweetness of ripened fruit because it hydrolysis starch into maltose during ripening of fruits. pH 4.0–5.5 is the optimum for the hydrolytic activity of β-amylase. β-Amylase has been used for various research and industrial applications. It is utilized for structural studies of starch and glycogen molecules produced by different methods. It is used in the brewing and

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distilling industry. It is also applied for producing high-maltose syrups (Sivaramakrishnan, Gangadharan, Nampoothiri, Soccol, & Pandey, 2006).

2.3 Glucoamylase Glucoamylase (EC 3.2.1.3) breaks α (1-4)glycosidic bonds as well as α(1-6) glycosidic bonds in the nonreducing end of amylose and amylopectin, producing glucose. Its optimum pH is 3.0 and can be efficiently utilized in acidic environments (Sivaramakrishnan et al., 2006).

3. PRODUCTION METHODS Submerged fermentations (SmF) and solid-state fermentations (SSF) are the two methods widely employed for the production of amylases. The solid-state fermentation is a new method, whereas SmF is a traditional method for enzyme production from microorganisms which has been used for a longer period of time. In SmF, free-flowing liquid substrates like molasses and broths are used. The end products of the fermentation are liberated into the fermentation broth. Substrate utilization is very rapid in SmF; henceforth, substrate must be provided continuously for this fermentation process. This technique is well suited for the extraction of secondary metabolites from bacteria because it requires high moisture content for their growth (Couto & Sanroma´n, 2006). SmF has several advantages in which genetically modified organisms are grown well compared to SSF and media sterilization, purification, and recovery of the end products. Further, the control of process parameters such as pH, temperature, moisture, oxygen transfer, and aeration can be done easily (Kunamneni, Permaul, & Singh, 2005). Solid-state fermentation (SSF) is suitable for the less moisture content required microorganisms. In SSF, nutrient-rich waste materials such as bran, bagasse, and paper pulp can be used as substrate for the microorganisms and they are consumed very slowly and constantly. Hence, there is no need to supply the substrate for longer time (Kunamneni et al., 2005). Major advantages of SSF are easy to handle, recovery of higher concentration of products, and generation of lesser effluent (Couto & Sanroma´n, 2006). Therefore, SSF is considered as a promising method for commercial enzyme production. α-Amylase production by SmF and solid-state fermentation techniques has been examined for fungal species. The results showed that

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SSF was well suited for developing countries due to cost-effective production process (Kunamneni et al., 2005).

4. PRODUCTION OF AMYLASES Despite several microorganisms can produce amylases, it is very difficult to find a most suitable strain for this enzyme production. Some microorganisms produce more than one enzyme, for e.g., α-amylase and glucoamylase. For example, Aspergillus niger can produce as many as 19 enzymes, whereas α-amylase can be produced in reasonably good titers by as many as 28 strains (Pandey, 1992). pH, temperature, nutrient supplementation, aeration, and the control of contamination during fermentation are the very crucial parameters necessary for the growth of the microorganisms in fermentation for large-scale production.

4.1 Production of α-Amylase Major source for the α-amylase production is Bacillus species, which can either be produced by solid substrate fermentation (SSF) (Babu & Satyanarayana, 1995; Krishna & Chandrasekaran, 1996; Ramesh & Lonsane, 1987) or by SmF (Kellly, Bloton, & Fogarty, 1997; Omidji, Amund, Braimoh, & IIori, 1997). The SSF is considered as a most economic technique in enzyme production as well as for starch hydrolysis. Cheese whey, corn steep liquor, and soya bean meal are the cheap sources for the industrial production of α-amylase (Bajapai, Gera, & Bajapai, 1992; Omidji et al., 1997). Marine bacteria produce α-amylases with specific features such as pH and thermo stability, salt tolerance, and cold activity. For example, deep sea bacterium Nocardiopsis sp. 7326 (Zhang & Zeng, 2008) and marine Wangia sp. C52 (Liu et al., 2011) produced cold-adapted α-amylase, pH stable α-amylase was derived from marine bacterium Bacillus subtilis S8-1 (Kalpana & Pandian, 2014) and Bacillus sp. ALSHL3 (Vidilaseris et al., 2009), and Zunongwangia profunda (AmyZ2) (Wu, Qin, Cheng, & Liu, 2014) have potential to produce salt-tolerant α-amylase. Alkalophilic amylase produced by sponge-associated marine bacterium Halobacterium salinarum MMD047 had optimum activity at 40°C and pH 9.0 (Shanmughapriya et al., 2009). The impact of calcium ion and EDTA on α-amylase production was evaluated and results revealed that calcium ions stimulated the activity as well as it was very essential for the thermostability of the enzyme whereas EDTA decreased the activity of α-amylase

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(Chakraborty et al., 2011; Vieille & Zeikus, 2001). In contrast to this, calcium ions neither stimulate nor decrease the activity of α-amylase in Streptomyces strain A3 and indicated the calcium independency of this strain (Chakraborty, Raut, Khopade, Mahadik, & Kokare, 2012). It was also reported that the activity of calcium-dependent α-amylase is reduced in the presence of EDTA (Kiran & Chandra, 2008), whereas calciumindependent enzyme retain their activity even in the presence of EDTA (Chakraborty et al., 2012). Chakraborty et al. (2014) isolated a novel 45 kDa α-amylase from marine Nocardiopsis sp. and immobilized it in gellan gum. They found 74.76  1.32% to 87.64  1.52% entrapment efficiency of this immobilized enzyme. A novel 66 kDa α-amylase was isolated from the marine Streptomyces sp. by Chakraborty, Khopade, Kokare, Mahadik, and Chopade (2009). The specific activity was 113.64 U/mg protein and almost 50% of this enzyme activity was retained at 85°C and the activity was not inhibited by the presence of commercially available detergent and oxidizing agents. Surfactants, oxidant, and detergent stable 66 kDa α-amylase were isolated from marine haloalkaliphilic Saccharopolyspora sp. A9. The enzyme was active in a wide range of NaCl concentration with maximum activity at 11% NaCl (Chakraborty et al., 2011). Marine actinomycetes isolated from South Indian coastal region produced 6.48 U/mL of amylase (Selvam, Vishnupriya, & Subash Chandra Bose, 2011). While comparing with this result, Streptomyces sp. NIOT-VKKMA02 isolated from Andaman & Nicobar Islands produced 13.27 U/mL of amylase enzyme, which is twofold increases to that of previous report (Meena, Anbu Rajan, Vinithkumar, & Kirubagaran, 2013). Mohapatra, Banerjee, and Bapuji (1998) isolated α-amylase from Mucor sp. and found that the optimum activity at pH 5.0 and temperature 60°C. It has also been reported that the activity was not inhibited in the presence of 3% NaCl, 10 mM Ca2+, and 25 mM Mg2+, whereas EDTA strongly inhibited the activity. Starch is considered as the major substrate for yeast cells and their primary, secondary metabolites productions because it is very cheap and easily available raw material in the world. In recent years, amylase-producing yeasts has been utilized drastically because of its potential role in the ethanol and single-cell protein production from starch (Chi, Liu, Ji, & Meng, 2003; Gupta et al., 2003).

4.2 Production of β-Amylase Usually plants are the major source of β-amylases. Henceforth, not much work has been done on the microbial β-amylase. Despite some

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microorganisms have the potential to produce β-amylase like Pseudomonas sp., Bacillus cereus, Bacillus polymyxa, Bacillus megaterium, Streptomyces sp., and Rhizopus japonicus (Crueger & Crueger, 1989; Fogarty & Kellly., 1990). β-Amylase production from starch waste by B. megaterium B6 mutant UN12 was compared in the SmF and SSF. Arum and wheat bran gave the highest yields (Ray, Jana, & Nanada, 1997).

4.3 Production of Glucoamylase Glucoamylase production from A. niger in solid cultures has been widely reported (Pandey, 1992; Selvakumar, Ashakumary, & Pandey, 1998). Agroindustrial residues such as wheat bran, rice bran, rice husk, gram flour, wheat flour, corn flour, tea waste, and copra waste were used (Pandey, 1992; Pandey & Radhakrishnan, 1990, 1993; Selvakumar et al., 1998). Marine yeast Aureobasidium pullulans N13d (Li, Chi, Wang, & Ma, 2007) produces 58.5 U/mg protein within 56 h of fermentation. The results revealed that crude glucoamylase actively digests potato starch granules, but poorly hydrolyzes raw corn and sweet potato starch. During hydrolysis only glucose molecules are released from starch, indicating that glucoamylase can effectively break both α-1,4 and α-1,6 linkages of starch (Li, Chi, Duan, et al., 2007; Li, Chi, Wang, et al., 2007).

5. PURIFICATION OF AMYLASES High purity amylases have been only used in pharmaceutical and clinical sectors as well as for studying structure–function relationships and biochemical properties of the enzymes (Gupta et al., 2003). Henceforth, it is very crucial for developing economic process for enzyme purification to obtain pure enzymes with maximum specific activity. Purification of amylases has been carried out in the following steps. Supernatant is obtained by centrifugation then it is concentrated by ultra filtration. The crude amylase can be precipitated and concentrated by ammonium sulfate precipitation or organic solvents such as ethanol in the cold. The precipitated sample is subjected to dialysis against water or a buffer for further concentration (Shih & Labbe, 1995) and followed by the chromatographic techniques such as ion exchange, gel filtration, and affinity. The number of steps used for purification of enzymes will depend on the extent of purity (Gangadharan, Sivaramakrishnan, Nampoothiri, & Pandey, 2006; Kundu & Das, 1970; Moreira et al., 1999; Sivaramakrishnan, Gangadharan, Nampoothiri,

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Soccol, & Pandey, 2007). Combination of filtration and centrifugation is used for obtaining extracellular enzyme from the fermented mass. For intracellular enzymes, raw corn starch may be added followed by filtration and subsequent steps. Enzyme produced by Aspergillus falvus var. columnaris was precipitated, dialyzed, and then subjected to column chromatography for purification (Shih & Labbe, 1995). For purification of amylase from Preussia minim, the sample was precipitated by trichloroacetic acid/acetone and eluted through Sephadex G-200 gel filtration column, and the resulting fractions were pooled and applied to a DEAE–Sepharose ion-exchange column (Erdal & Taskin, 2010). Ammonium sulfate (80%) precipitation, TSK Toyopeal column chromatography, ultrafiltration, dialysis, and SP Sepharose column chromatography steps were used for the purification of α-amylase from mutant Bacillus (Ballschmiter, F€ utterer, & Liebl, 2006). For thermostable amylase purification, the cell extract obtained after centrifugation is subjected to high temperatures for denaturing thermolabile proteins. This step was followed by anion exchange chromatography for purification of α-amylase from Thermotoga maritima MSB8. Then SDSPAGE and size exclusion chromatography were used for checking the purity of the enzyme as well as for the determination of its molecular weight (Iefuji, Chino, Kato, & Iimura, 1996).

6. CHARACTERIZATION OF AMYLASES After purification, characterization is carried out for enzymes. SDS-PAGE is mostly used for characterization of purified enzyme along with molecular markers like BSA (67 kDa) and ovalbumin (43 kDa). Then the gel is stained with either Coomassie Brilliant Blue (Paquet, Croux, Goma, & Soucaille, 1991) or silver nitrate (Mukesh Kumar, Jayanthisiddhuraj, Monica Devi, Bala Kumaran, & Kalaichelvan, 2012) and visualized. Purity and homogeneity of the enzyme is also observed in the PAGE. Molecular mass of α-amylase produced by Bacillus MNJ23 and mutant B. subtilis was characterized by SDS-PAGE (12% and 10%, respectively) (Ballschmiter et al., 2006; Singhania, Patel, Soccol, & Pandey, 2009). SDSPAGE in a Phast Gel gradient (10–15%) was used for the characterization of enzyme produced by Clostridium acetobutylicum ATCC 824 (Mukesh Kumar et al., 2012). Size exclusion chromatography was also used for determining the molecular mass of the enzyme (Satoh, Uchimura, Kudo, & Komagata, 1997).

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Intracellular α-amylase of Streptococcus bovis 148 showed a single band on a SDS-PAGE gel which indicated the purification to homogeneity. The molecular mass was found to be 57,000 Da, near the 55,000 Da protein obtained with the gel filtration method, confirming that the intracellular α-amylase exists in a monomeric form (Melasniemi, 1987). Dong, Vieille, Savchenko, and Zeikus (1997) used Sephacryl S200 gel filtration chromatography for determining molecular weight of α-amylase produced from Pyrococcus furiosus. In addition to SDS-PAGE, isoelectric focusing is also used for enzyme characterization. The isoelectric point of the desired enzyme has been determined by using protein markers of broad range of pI values (Mukesh Kumar et al., 2012). α-Amylase produced by Clostridium thermohydrosulfuricum E 101-69 was ultrafiltered for concentration of crude enzyme sample and then it was subjected to isoelectric focusing with pI range of 3.5–5.0 (Nielsen & Borchert, 2000). pI 3–10 as standards was used for determining isoelectric point of C. acetobutylicum ATCC 824 α-amylase (Mukesh Kumar et al., 2012).

7. MOLECULAR BIOLOGY OF AMYLASES Recombinant DNA technology offers great features for cloning of amylase-producing strains, particularly α-amylase and glucoamylase. The main aim of gene cloning is the overexpression of thermostable enzymes, maximum productivity, and expression of the two enzymes simultaneously by the same strain. Much work has been carried out on the cloning of α-amylase genes either in Escherichia coli or in Saccharomyces cerevisiae. Like α-amylase, the number of glucoamylase-encoding genes also cloned in E. coli. Only very few reports are available on β-amylase expression in E. coli, usually from Bacillus sp. and only one report from Thermoanaerobacterium.

8. DETERMINATION OF ENZYME ACTIVITY Reducing sugars released as a result of starch hydrolysis by amylase enzyme is measured for determining the enzyme activity. Reading the absorbance of starch–iodine complex is another method for measuring enzyme activity. The following methods are usually used for enzyme assay.

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8.1 Dinitrosalicylic Acid Method In this method, enzyme solution is added to the aliquots of the substrate stock solution and incubated at 50°C for 10 min followed by the addition of dinitrosalicylic acid (DNS) method and the mixture is incubated in a boiling water bath for 5 min. After cooling to room temperature, the absorbance of the supernatant is measured at 540 nm. The activity is calculated by subtracting the A540 values of substrate and enzyme blanks from the A540 value of the analyzed sample (Feller, Le Bussy, & Gerday, 1998; Gusakov, Kondratyeva, & Sinitsyn, 2011; Kobayashi et al., 1992). DNS method was used for measuring alkalophilic α-amylase activity of Bacillus strain GM8901 (Gusakov et al., 2011).

8.2 Nelson–Somogyi Method In this method, enzyme solution and substrate stock solutions are heated at 50°C for 5 min separately. Then enzyme solution is added to the substrate stock solution and this mixture is incubated at 50°C for 10 min. After 10 min, Somogyi copper reagent is added to this mixture to terminate the reaction. This is followed by incubation of this mixture in a boiling water bath for 40 min and cooled to room temperature. Then Nelson arsenomolybdate reagent is added and incubated at room temperature for 10 min. Finally, water is added and absorbance of supernatant is read at 610 nm (Kobayashi et al., 1992). Haloalkaliphilic α-amylase of archaebacterium Natronococcus sp. Strain Ah-36 was subjected to NS method for determining enzyme activity (El-Safey & Ammar, 2004).

8.3 Determination of Activity Using Iodine This method is used for enzyme assay based on the principle that the formation of blue colored complex by the reaction of starch with iodine. On hydrolysis of starch by amylase enzyme this complex is changed into reddish brown color. Then the absorbance is read after the termination of enzyme substrate reaction. This gives a measure of the extent of hydrolysis of starch by α-amylase (Gupta et al., 2003).

8.4 Dextrinizing Activity In this enzyme assay, the crude enzyme is incubated at 92°C with soluble starch for 10 min. Then the reaction mixture is kept in ice to terminate the reaction. Iodine is added to this reaction mixture to form a colored

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complex with starch and then water is added to this complex to dilute the color to a measurable range that can be read at 600 nm (Haki & Rakshit, 2003).

8.5 Indian Pharmacopeia Method This method is used for determining the enzyme activity by measuring the amount of starch (grams) hydrolyzed by enzyme of a given volume. In this assay different concentration of enzyme solution is incubated with starch at 40°C for 1 h. After incubation, iodine is added to this mixture to produce colored complex (Gupta et al., 2003).

8.6 Reduction in Viscosity of Starch Suspension This method is mostly used in baking industry to check the quality of flour. Falling number (FN) method and Amylograph/Farinograph test are used for determining enzyme activity based on decreasing viscosity of starch suspension. In FN method, enzyme substrate preparations are measured at 100°C. Usually malted flour has a FN of around 400. Amylograph test is used to test the relationship between the enzyme activity and peak starch slurry viscosity. The lesser viscosity indicates the higher activity of the enzymes. Optimal Brabender units for bread baking flour ranges from 400 to 600 units (Gupta et al., 2003).

9. INDUSTRIAL APPLICATIONS AMYLASES In recent years interest in amylase production has been increased drastically due to its starch hydrolyzing properties and utilization of easily available and low cost raw material. It has many potential and widely used applications on the industrial front. In various industrial sectors, chemical methods used for the hydrolysis have been replaced by enzymes and it makes the process easier and eco-friendly.

9.1 Enzymatic Hydrolysis of Starch for Fructose and Glucose Production Fructose and glucose syrups are produced by the hydrolysis of starch by amylase (Van Der Maarel, Van Der Veen, Uitdehaag, Leemhuis, & Dijkhuizen, 2002). This process consists of three steps. Gelatinization is the first step followed by liquefaction and saccharification. In gelatinization starch granules are dissolved in water to produce a viscous starch suspension. Then

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amylase is added into the water to initiate hydrolysis. Starch is partially hydrolyzed by amylase and form short chain of dextrins resulting in reduction of the starch suspension. This process is called liquefaction. Release of glucose from starch by further hydrolysis by glucoamylase is called saccharification. α-1,4-Glycosidic linkages of nonreducing terminal ends are hydrolyzed by glucoamylase cleaves from the nonreducing terminal ends of starch, hence, it is called as an exoamylase. Pullulanase activity along with glucoamylase produces high-glucose syrup. Glucose isomerase converts the high-glucose syrup into high-fructose syrup by catalyzing the isomerization. The fructose syrup is usually used as a sweetener, especially in the beverage industry. Traditionally, acid hydrolysis is used for the production of glucose and fructose syrup. This process had many drawbacks. Due to the acidic nature of this process, it required corrosion-resistant equipment and high temperatures would inactivate the thermolabile enzymes. Henceforth, enzymatic hydrolysis is preferred for glucose and fructose syrup production. The α-amylase used in the liquefaction step can be derived from different microbial sources, for example, Bacillus amyloliquefaciens, Bacillus licheniformis, Bacillus stearothermophilus, and P. furiosus are mostly used for α-amylase production. Hydrolysis of starch at a high temperature can be carried out with the help of thermostable amylases (Gupta et al., 2003; Kulp, Ponte, & D’Appolonia, 1981).

9.2 Bakery Industry For starch hydrolysis of dough, it is supplemented with α-amylase followed by yeast fermentation. Undesirable changes such as increase of crumb firmness, loss of crispness, decrease in moisture content, and loss of bread flavor occur during storage of baked bread. This together is called staling. α-Amylase acts as an antistaling agent to increase the shelf life of baked breads (Gupta et al., 2003; Kulp et al., 1981). However, a slight overdose of this enzyme produces gumminess of the bread (Chi et al., 2009). To prevent this, pullulanase is applied along with amylase for breakdown of gumminess responsible compounds (Kulp et al., 1981).

9.3 Other Industrial Applications Enzymes are employed in detergent industry due its eco-friendly nature and have the potential to work at mild conditions. The potential of α-amylase to work at low temperature and alkaline pH makes it extensively used in detergents. Calcium dependency and oxidant sensitivity are the major drawbacks

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of α-amylase. The oxidant resistance potential of amylase is increased by replacing the oxidant-sensitive amino acid met by leu in B. licheniformis amylase (Gupta et al., 2003). Starch is a preferred sizing agent due to its easy availability, low cost and can be easily removed from the fabric. α-Amylase is used for hydrolyzing starch into water-soluble components during desizing process where that can be removed by washing. The enzyme acts only on the starch molecules, it does not affect the fibers (Gupta et al., 2003). For protection of papers from stains in processing it is also coated with sizing agents. Starch is the most preferred sizing agent in paper industry. Starch is partially hydrolyzed by α-amylase and reduces the viscosity in a batch or a continuous process and makes it a suitable sizing agent for paper industry (Gupta et al., 2003). Mostly starch is used as a starting material for biofuel production due its low cost and easy availability. First step is liquefaction where starch is converted into viscous suspension. This suspension is subjected to saccharification for the production of fermentable sugars through hydrolysis by α-amylase. Finally, ethanol is produced from these fermentable sugars by yeast fermentation. S. cerevisiae and amylolytic yeast Saccharomyces fibuligera are fused for protoplasmic fusion to generate a new yeast strain to yield biofuel from starch to exclude saccharification process (Chi et al., 2009).

10. CONCLUSION The use of enzyme for various industrial needs has been increased recently due to their biodegradability and can be obtained from biological sources. By-products and waste products of other processes can be utilized for amylase production from microorganisms. The enzyme has many industrial applications which include production of fructose syrup, eco-friendly detergents, and baked products. It is also used as alternative for fossil fuels depletion by producing biofuel such as ethanol from starch. From these applications, it can be concluded that it is a promising agent to replace the conventional chemical process usually employed in the industries.

ACKNOWLEDGMENT Dr. J.S. is grateful to University Grants Commission—Dr. D.S. Kothari acknowledges the Postdoctoral Fellowship for their financial support.

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CHAPTER NINE

Enzyme Immobilization: An Overview on Methods, Support Material, and Applications of Immobilized Enzymes V.L. Sirisha*,1, Ankita Jain*,†, Amita Jain*,{ *UM-DAE Centre for Excellence in Basic Sciences, University of Mumbai, Mumbai, India † University of Rajasthan, Jaipur, India { D.Y. Patil University, Navi Mumbai, India 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 1.1 Adsorption/Carrier-Binding Method 1.2 Covalent Binding 1.3 Affinity Immobilization 1.4 Entrapment 1.5 Ionic Binding 1.6 Metal-Linked Immobilization 2. Materials Used for Fabrication of Immobilization Supports 2.1 Inorganic Material as Supports 2.2 Organic Supports 3. Applications of Immobilized Enzymes 3.1 Immobilized Enzymes as Biosensors 3.2 Immobilized Enzymes in Medicine and Antibiotics 3.3 Immobilized Enzymes in Food and Dairy Industry 3.4 Immobilized Enzyme in Bioremediation/Waste Water Treatment 3.5 Biodiesel Production 3.6 Immobilized Enzymes in Biotech Cleaning 3.7 Carbon Capture 4. Conclusions References

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Abstract Immobilized enzymes can be used in a wide range of processes. In recent years, a variety of new approaches have emerged for the immobilization of enzymes that have greater efficiency and wider usage. During the course of the last two decades, this area has rapidly expanded into a multidisciplinary field. This current study is a comprehensive review

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of a variety of literature produced on the different enzymes that have been immobilized on various supporting materials. These immobilized enzymes have a wide range of applications. These include applications in the sugar, fish, and wine industries, where they are used for removing organic compounds from waste water. This study also reviews their use in sophisticated biosensors for metabolite control and in situ measurements of environmental pollutants. Immobilized enzymes also find significant application in drug metabolism, biodiesel and antibiotic production, bioremediation, and the food industry. The widespread usage of immobilized enzymes is largely due to the fact that they are cheaper, environment friendly, and much easier to use when compared to equivalent technologies.

1. INTRODUCTION The process of biocatalysis has been immensely used in various sectors of biotechnology because of their high substrate specificity, ease of production, and green chemistry. Use of enzymes for large extent commercialization becomes limited depending on the costs, because those more expensive are not economical because of their low reusability factor. Moreover, the structural stability of some of the enzymes during any biochemical reaction is highly challenging. In order to overcome these limitations, immobilization of useful enzymes with functional efficiency and increased reproducibility is found to be more promising in spite of their expensiveness. Enzymes and whole cells can be used as immobilized biocatalysts (Kawaguti, Manrich, & Sato, 2006). Enzyme immobilization may be defined as confining the enzyme molecules to a solid matrix/support different from the one in which substrate or the products are present. This is achieved by attaching the enzymes to or within some suitable support material. It is important to note that the substrate molecules and the products formed should move freely in and out of the phase to which the enzymes are restrained. Various materials can be used as matrix or support system for enzyme immobilization, generally inert polymers and inorganic materials. The ideal carrier matrix should have the following properties: (i) to be economical, (ii) inertness, (iii) stability, (iv) physical strength, (v) ability to enhance enzyme specificity/activity, (vi) regenerability, (vii) ability to reduce product inhibition, and (viii) ability to prevent nonspecific adsorption and bacterial contamination (Singh, 2009). Most of the matrices have only some of the properties; hence, selection of the carrier matrix for enzyme immobilization must be chosen based on the properties and limitations of the matrices. The process of immobilization produces continuous economic operations, automation,

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high investment/capacity ratio, and recovery of product with greater purity (D’Souza, 1998). There are different methods of enzyme immobilization and various factors impact the immobilized enzymes performance. The different enzyme immobilization methods are grouped as follows and listed in Fig. 1: 1. Adsorption/carrier-binding method. 2. Covalent bonding/cross-linking. 3. Entrapment method. 4. Membrane confinement. In this chapter, the various immobilization methods along with latest development for each of them are discussed. Special emphasis is given to the potential modifications for each method and the support material employed for immobilization that may have significant consequences on future enzyme market.

1.1 Adsorption/Carrier-Binding Method In this method, the enzyme molecules adhere to the surface of the carrier matrix by a combination of hydrophobic interactions and the formation of various salt linkages per molecule of enzyme. The process involves the suspension of matrix with the enzyme or drying of the enzyme molecules on electrode surfaces. The most commonly used matrix materials are water soluble carriers like polysaccharide derivatives, glass, synthetic polymers, etc. (AlAdhami, Bryjak, Greb-Markiewicz, & Peczynska-Czoch, 2002; Cordeiro, Lenk, & Werner, 2011; Rosa, Cruz, Vidal, & Oliva, 2002; Wu & Lia, 2008). In this method, there will be a strong bond between the enzyme and the support matrix. However, some physiological conditions like high temperature or pH, substrate addition may weaken the bond. The adsorbed enzymes are usually resistant to proteolysis and aggregation because of their hydrophobic interaction with interfaces (Spahn & Minteer, 2008). The other commonly used matrix material is the coconut fibers because of their waterholding potential and enhanced cation exchange policy. Kaolin, another ecofriendly matrix upon acetylation, showed high enzyme retainability. Other matrix materials include micro/mesoporous substances having large surface area and functionalized thiol group, microcrystalline cellulose with irreversible-binding capacity, etc., have been used by various researchers (Brı´gida, Calado, Goncalves, & Coelho, 2010; Dey, Nagpal, & Banerjee, 2002; Herna´ndez et al., 2007; Huang et al., 2011; Karagulyan, Gasparyan, & Decker, 2008; Mitchell & Ramı´rez, 2011).

Fig. 1 Various methods of enzyme/cell immobilization.

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Because of presence of silanols, silanized molecular sieves facilitate enzyme immobilization by hydrogen bonding (Diaz & Balkus, 1996). For better immobilization, different chemical modifications of the present support matrices would be promising. Persson, Wehtje, and Adlercreut (2000) adsorbed lipase enzyme on polypropylene-based hydrophobic granules/Accurel EP-100 and studied the lipase water activity profiles. It was found that during biocatalysis, reducing the particle size of Accurel significantly increased the enantiomeric ratio and reaction rates (Sabbani, Hedenstr€ om, & Nordin, 2006). It was reported that when Yarrowia lipolytica lipase was immobilized on octyl-agarose and octadecyl-sepa beads supports by physical adsorption, resulted in greater stability, higher yields, better process control, and quite economical as compared to free lipase. This was mainly because of the hydrophobicity of octadecyl-sepa beads that increases the enzyme and support affinity (Cunha et al., 2008). When lipase from Candida rugosa adsorbed onto poly(3-hydroxybutyrate-co-hydroxyvalerate), expressed 94% activity even after treating for 4 h at 50°C and reusing till 12 cycles (CabreraPadilla et al., 2011). Mishra, Pithawala, and Bahadur (2011) reported increased pH stability and ability to retain 50% activity under dried condition of urease enzyme by adsorbing onto 1,4-butenediol diglycidyl etheractivated byssus threads. These reports clearly showed that eco-friendly support matrix materials found to be economical, biocompatible, biodegradable, long durable, efficient, and also prevents ethical issues (Popat et al., 2011).

1.2 Covalent Binding This method is mainly depends on the formation of covalent bond between the enzyme and the support material. Covalent bond formation between the enzyme and the matrix happens through the side chain amino acids like histidine, arginine, aspartic acid, etc. However, the reactivity depends on the presence of different functional groups such as carboxyl group, amino group, indole group, phenolic group, sulfhydryl group, thiol group, imidazole group, and hydroxyl group. By preventing the active site amino acid residues inactivation, efficient enzyme activities can be achieved. Widely used protective strategies are 1. Covalent binding of the enzyme with the matrix in the presence of substrate or a competitive inhibitor. 2. By formation of a reversible covalently linked enzyme–inhibitor complex.

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3. Covalent linkage of a chemically modified soluble enzyme to the matrix is obtained by incorporating new residues. 4. A zymogen precursor. Moreover, higher specific activity and stability with controlled protein orientation can be achieved when peptide-modified surfaces are used for enzyme linkages (Fu, Reinhold, & Woodbury, 2011). For example, high thermal stability of the enzyme was exhibited when it is covalently linked to cyanogen bromide (CNBr)-agarose and CNBr-Sepharose, having carbohydrate moiety and glutaraldehyde as spacer arm (Cunha et al., 2008; Hsieh, Liu, & Liao, 2000). Moreover, covalently binding of enzymes to modified silica gel carriers (by removing unreacted aldehyde groups with SBA-15 supports) has shown enhanced enzyme stability and also acts as hyperactive biocatalysts (Lee, Park, Yeo, & Kim, 2006; Szymans´ka, Bryjak, & Jarzebski, 2009). Moreover, covalent binding of enzymes with mesoporous silica and chitosan showed to increase the half-life and thermal stability of the enzymes (Hsieh et al., 2000; Ispas, Sokolov, & Andreescu, 2009). The use of nanodiametric supports for enzyme immobilization has brought a turning point in this field. It was reported that cross-linking of enzymes with electrospun nanofibers has showed greater residual activity because of increase porosity and surface area (Huang & Cheng, 2008; Kim, Jia, & Wang, 2006; Li, Chen, & Wu, 2007; Ren et al., 2006; Sakai et al., 2010; Wu, Yuan, & Sheng, 2005). Researchers in this field used attapulgite nanofibers as support material for covalent binding with alcohol dehydrogenase enzyme owing to its thermal tolerance and are available in different nanosizes (Zhao et al., 2010). Immobilization of enzymes on magnetic nanocluster by covalent binding with different orientations showed varied applications in pharmaceutical industries because of their enhanced operational stability, reusability, and longevity (Yusdy, Patel, Yap, & Wang, 2009). Maintenance of immobilized enzymes structural and functional properties is very important which can be played by a cross-linking agent. Glutaraldehyde is one such cross-linking agent, due to its solubility in aqueous solvents and can form stable inter- and intrasubunit covalent bonds, popularly used as bifunctional cross-linker.

1.3 Affinity Immobilization Affinity binding is the immobilization of enzyme to the support matrix by specific interactions. Affinity immobilization utilizes the enzyme specificity to its support under various physiological conditions. This can be achieved

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by following two methods. The first method is precoupling of the matrix to an affinity ligand for target enzyme. The other method consists of the enzyme modified or conjugated to another molecule which develops affinity toward the matrix (Nisha, Arun Karthick, & Gobi, 2012; Sardar, Roy, & Gupta, 2000). Affinity adsorbents have also been used for enzyme purification (Ho, Li, Lin, & Hsu, 2004). Enzymes immobilized on complex affinity support matrix such as agarose-linked multilayered concanavalin A and alkali stable chitosan-coated porous silica beads possess higher amounts of enzyme and promoted increased efficiency and stability (Sardar & Gupta, 2005; Shi, Tian, Dong, Bai, & Sun, 2003). The enzyme-binding capacity and reusability can be increased significantly by using bioaffinity layering method. This is mainly because of the presence of noncovalent forces like van der Waals forces, columbic, hydrogen bonding, etc. (Haider & Husain, 2008; Sardar & Gupta, 2005).

1.4 Entrapment The entrapment method is based on the caging of the enzyme within a polymeric network by covalent or noncovalent bonds that allow the passage of substrate and products but retain the enzyme (Singh, 2009). In this method the enzyme is not bound to the support matrix unlike other methods which are described earlier. The caging of enzyme can be achieved by any of the following strategies: (1) by inclusion if enzyme within a highly cross-linked polymer matrix, (2) by enzyme dissolution in a nonaqueous phase, or (3) by separating enzyme from a bulk solution by using a semipermeable microcapsule. There are various methods of enzymes entrapment like fiber entrapping (Dinelli, 1972), gel entrapping (Bernfeld & Wan, 1963), microencapsulation (Wadiack & Carbonell, 1975), etc. Encapsulation with alginate–gelatin–calcium hybrid carriers has been reported to be efficient, as it increased mechanical stability and also prevented enzyme leakage (Shen et al., 2011). In C. rugosa, when the lipase enzyme was entrapped in chitosan, it showed enhanced enzyme activity and entrapment efficiency. It also prevented friability and leaching. This is mainly because the support matrix is biocompatible and nontoxic; receptive to chemical modifications because of its hydrophilic nature it has high affinity toward proteins (Betigeri & Neau, 2002). Chen, Kuan, et al. (2011) reported that simultaneous entrapment of lipase and magnetic nanoparticles with biomimetic silica significantly increased the enzyme activity under various silane additives. It has also been reported that lipases when entrapped with ĸ-carrageenan showed

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high tolerance to organic solvents and also found to be highly thermostable (Jegannathan, Jun-Yee, Chan, & Ravindra, 2010; T€ umt€ urk, Karaca, Demirel, & Sahin, 2007). Use of nanostructured supports like electrospun nanofibers, pristine materials for entrapment of enzymes have found to have varied applications in the fields of biomedicine, biofuel, chemistry, biosensors, etc. (Dai & Xia, 2006; Kim et al., 2006; Wang, Wan, Liu, Huang, & Xu, 2009; Wen, Nallathambi, Chakraborty, & Barton, 2011).

1.5 Ionic Binding The ionic binding method relies on the ionic binding of the enzyme protein to water-insoluble carriers containing ion-exchange residues. This type of noncovalent immobilization can be reversed by altering the ionic strength, polarity, and temperature. This principle is similar to protein– ligand interactions principles used in chromatography (Guisa´n et al., 1997). Support materials such as polysaccharides and synthetic polymers having ion-exchange centers are generally used in this type of enzyme immobilization. There are advantages of this method like the binding of the enzyme with the carrier is much simpler and the conditions used are milder than covalent binding. Moreover, ionic binding causes slight modifications in the conformation and active site of the enzyme, hence leading to high enzyme activity in most of the times. However, enzyme leakage from the carrier may happen, when substrate solution of high ionic strength or varied pH solutions are used. This is because of weak bonding between the enzyme proteins and the carrier molecules. In case of ionic binding the bond between enzyme and carrier linkages are much stronger than physical adsorption. However, ionic binding is weaker than covalent binding.

1.6 Metal-Linked Immobilization This method involves the precipitation of metal salts on the support matrix surface. These metals have the ability to bind to the nucleophilic groups of the carrier. The precipitation of the metal ion on the support matrix can be achieved by heating. The enzyme immobilized by this method shows relatively 30–80% higher enzyme activity. It is a simple, easy, and reversible process. By decreasing the pH of the solution, the enzyme and carrier molecule can be separated (Yucel, 2011). Moreover, both enzyme and matrix can be regenerated by this method.

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2. MATERIALS USED FOR FABRICATION OF IMMOBILIZATION SUPPORTS The materials used for immobilization of enzymes, called carrier or support matrices. The characteristics of matrix are of utmost importance in determining the success and result of immobilizing enzyme. Ideal support properties include the following properties: (i) It should be of low cost and eco-friendly, reducing the economic impact of the process. (ii) It should be totally inert after immobilization and not blocking the desired reaction. (iii) It should have thermal and mechanical resistance, allowing the immobilized enzyme to be used under various operational conditions. (iv) It should be highly stable. (v) It should have high regenerability after the useful lifetime of the immobilized enzyme. (vi) It should enhance the enzyme specificity. (vii) It should be able to pack large amount of enzyme. As such, the porosity plays a major role. Large pore size will cause a notable fall in the surface area, while small pore size will barely exclude the protein. So, diameter of the pore should be in appropriate range (Cao, 2006; Hartmann & Kostrov, 2013; Magner, 2013; Tran & Balkus, 2011; Weetall, 1976). (viii) To prevent undesired protein adsorption and denaturation, hydrophobicity of the support matrix surface should be minimized (Cao, 2006; Zhou & Hartmann, 2012). Therefore to increase the immobilized enzyme catalytic features, the support matrix should have required optimal environment (Rodrigues, Ortiz, BerenguerMurcia, Torres, & Ferna´ndez-Lafuente, 2013). (ix) The support should be able to shift in the pH optimum for enzyme action to the desired value for the process. (x) It should have antimicrobial and nonspecific adsorption properties. (xi) Nevertheless, most of the support matrices possess few of the above mentioned properties. So, care should be taken in selecting the appropriate support material keeping in mind their pros and cons of their properties. On the basis of their chemical composition the support matrices are classified under two main categories namely: (a) inorganic support material

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and (b) organic support material (organic supports are further subdivided into natural and synthetic supports).

2.1 Inorganic Material as Supports They include glass, silica gel, alumina, metal oxides, zirconia, and many other silica-based materials are considered as material of choice and are widely used as they have thermal and mechanical resistance. They also offer microbial resistance as inorganic supports does not serve as substrate for bacterial/fungal growth. The characteristic feature of inorganic support is they provide rigidity and porosity. Furthermore, they ensure invariance of pore diameter/pore volume which ensures fixed volume and shape to the support (Hartmann & Kostrov, 2013). The various inorganic support matrices, their properties, and the enzymes immobilized on this support are listed in Table 1. 2.1.1 Silica Silica-based inorganic support like silicon dioxide and silicon tetraoxide are generally used for enzyme immobilization. They exist in 3D polymer. Where SiO4 are rigid but SiO2 are flexible. Coexistence of both hydrophilic and hydrophobic sites gives it complex adsorptive properties. Another example of siliceous microporous support is zeolite which is popular for giving high surface area for immobilizing protein. Silica carriers are chemically inert, and they need modification and proper activation. So they are modified usually by treating aminoalkyl triethoxysilanes to introduce amino group, and further they are activated by variety of methods for enzyme immobilization. For example, penicillin G amidase was first linked with dextran and was then immobilized onto amino-activated silica gel, showed increased thermal stability (Burteau, Burton, & Crichton, 1989). Similarly, when lignin peroxidase and horseradish peroxidase (HRP) were immobilized on activated silica support, the immobilized enzymes found to remove chlorolignins effectively from eucalyptus kraft effluent (Dezott et al., 1995). It was also reported that immobilization of α-amylase on silica nanoparticles improves the cleaning performance of detergents (Soleimani et al., 2011). It was also reported that the enzyme–carrier bonding can be strengthened by surface modifications of silica, like amination of hydroxyl and reactive siloxane groups, addition of methyl or polyvinyl alcohol groups, etc. (Pogorilyi, Siletskaya, Goncharik, Kozhara, & Zub,

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Table 1 Inorganic Material, Their Properties, and Immobilized Enzymes Inorganic Enzyme Material Properties Immobilized References

Liu, Hu, and Deng (1997), Xing, Li, Tian, and Ye (2000), and Chang and Chu (2007)

Large specific surface Zeolite (molecular area of the zeolite substrate resulted in high sieves) enzyme loading

Glucose oxidase (GOx), α-chymotrypsin, lysozyme

Ceramics

Both macro- and micropores efficient in lowering diffusion rate and increasing specific surface area

Candida antarctica Magnan, Catarino, lipase Paolucci-Jeanjean, Preziozi-Belloy, and Belleville (2004)

Celite

Inexpensive, low polarity, large adhesion area, chemically inert, resistance against pH, temperature, urea, detergents, and organic solvents

Lipase, polyphenol oxidase, β-galactosidase

Khan, Akhtar, and Husain (2006), Liu, Lin, Chen, and Chang (2009), and Ansari and Husain (2012)

Silica

Nanosized structures, high surface area, ordered arrangement, high stability to chemical and mechanical forces, high mechanical strength

Lignin peroxidase, horseradish peroxidase, α-amylase

Dezott, InnocentiniMei, and Dura´n (1995) and Soleimani, Khani, and Najafzadeh (2011)

Glass

Highly viscous liquid

α-Amylase, nitrite reductase

Kahraman, Bayramoglu, Kayaman-Apohan, and G€ ung€ or (2007) and Rosa et al. (2002)

Activated carbon

Large contact sites, high Acid protease, surface area acidic lipase

Charcoal

Excellent adsorbent, minimum fine particulate matter release

Kumar, Perinbam, Kamatchi, Nagesh, and Sekaran (2010) and Ramani et al. (2012)

Papain, Dutta, Bhattacharyya, amyloglucosidase De, Ray, and Basu (2009) and Rani, Das, and Satyanarayana (2000)

Adapted from Datta, S., Christena, L. R., & Rajaram, Y. R. S. (2013). Enzyme immobilization: An overview on techniques and support materials. 3 Biotech, 3, 1–9.

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2007; Rao, Kembhavi, & Pant, 2000; Shioji, Hanada, Hayashi, Tokami, & Yamamoto, 2003). 2.1.2 Ceramics Ceramics are another kind of inorganic support which are solid, insoluble, inorganic, nonmetallic, and are obtained by cooking plastic materials. Metal oxides such as Al2O3, TiO2, ZrO2, and SnO2 are referred as ceramics (Bertuoli, Piazza, Scienza, & Zattera, 2014; Dimitrov et al., 2014; Jean, Sciamanna, Demuynck, Cambier, & Gonon, 2014; Osma, TocaHerrera, & Rodrı´guez-Couto, 2010; Ponto´n et al., 2014). Because of their porous nature, ceramics are greatly used for enzyme immobilization. It was reported that the immobilization of lipase enzyme from Candida antarctica on ceramic membrane helps in carrying out hydrolytic and synthetic reactions by restraining feedback inhibition (Magnan et al., 2004). Ceramic foams were also found to be effective in increasing the specific surface area and in decreasing diffusion rates (Magnan et al., 2004). 2.1.3 Glass It is another most commonly used inorganic support system. It is highly viscous in nature and has been used to immobilize various enzymes like α-amylase. In this process, phthaloyl chloride-containing amino group functionalized glass beads were found to be strong and sustainable. Similarly, urease, when immobilized on glass pH-electrodes, served as biosensor for monitoring urea in blood samples at levels as low as 52 mg/mL (Sahney, Puri, & Anand, 2005). 2.1.4 Charcoal Charcoal as support system, for immobilizing enzymes like amyloglucosidase without any cross-linking agents for starch hydrolysis, has been reported and found to have 90% catalytic activity (Rani et al., 2000). Charcoal was also found to be an efficient adsorbent with high adsorptive capacity and less particulate matter release (Kibarer & Akovali, 1996). 2.1.5 Activated Carbon It was reported that both natural and HCl-modified activated carbon was found to be a good support system for enzyme immobilization (Alkan et al., 2009). Recently, acid protease and acidic lipases were immobilized on mesoporous-activated carbon particles and showed significantly higher

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catalytic efficiency even after reusing for 21 cycles (Kumar et al., 2010; Ramani et al., 2012). Daoud, Kaddour, and Sadoun (2010) reported that efficient enzyme immobilization can be obtained by using activated carbon with high surface area (600–1000 m2/g
1) and pore volume (300–1000 A˚).

2.2 Organic Supports 2.2.1 Natural Polymers A wide variety of natural polymers, mainly water-insoluble polysaccharides like collagen, chitosan, carrageenans, alginate, cellulose, starch, agarose, etc., have been used as support matrix for enzyme immobilization. The characteristic features of these polymers that make them good support system include, their ability to form inert gels, due to their chemical structure which can be activated easily, they can bind to proteins and enzymes in a reversible and irreversible way, they are available in large quantities, inexpensive, and they show high thermal and mechanical resistance by cross-linking with bifunctional reagents. The various organic supports and their immobilized enzymes are listed in Table 2. 2.2.1.1 Alginate

It is a sulfated polysaccharide, obtained from brown algal cell wall. Immobilization of enzymes with the salts of alginate like xanthan–alginate beads, alginate–polyacrylamide gels, and calcium alginate beads give increased enzyme activity and reusability. As alginate cross-links with divalent ions and also glutaraldehyde, it improves enzyme stability (Elcin, 1995; FloresMaltos et al., 2011). 2.2.1.2 Chitosan

It is a polysaccharide which is been used as a support matrix (Kapoor & Kuhad, 2007; Vaillant et al., 2000). It has various desirable properties that make it a suitable carrier system. These include mechanical stability, rigidity, hydrophilicity, active reactive groups that can interact with enzymes directly and affinity to bind proteins. It was reported that the enzymes coated with chitosan has less leaching effect compared to alginate because of its ionic and physical interactions with the enzyme (Betigeri & Neau, 2002). It was reported that a wet composite of chitosan and clay which has both hydroxyl and amino groups were found to be efficient for enzyme immobilization along with enhanced hydrophilicity and high porosity. It was observed in Bacillus circulans, the chitin-binding

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Table 2 Organic Support System, Their Properties, and Immobilized Enzymes Natural Polymers as Enzyme Support Properties Immobilized References

Elcin (1995) and Flores-Maltos et al. (2011)

Alginate

Used as xanthan– alginate beads, alginate– polyacrylamide gels, and calcium alginate beads Can be reused, improves the stability of enzymes

Chitosan and chitin

Used in combination with alginate Less leaching effect, more reliable for enzyme trapping, in form of beads can entrap more enzymes

Collagen

Retain significant Tannase, catalase activity after many cycles of reuse

Katwa, Ramakrishna, and Rao (1981) and Chen, Song, Liao, and Shi (2011)

Carrageenan (linear sulfated polysaccharide)

Improve stability, Lipase, as pseudoplastic in α-galactosidase nature it helps it to thin under shear stress and recover its viscosity once stress is removed Cheap, durable, better entrapment

T€ umt€ urk et al. (2007), Jegannathan et al. (2010), Rao, Prakasham, Rao, and Yadav (2008), and Girigowda and Mulimani (2006)

Gelatin (hydrocolloid High adsorption material, high in ability, promote loading efficiency amino acid)

D-Hydantoinase

Betigeri and Neau (2002) and Chang and Juang (2007)

Shen et al. (2011)

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Table 2 Organic Support System, Their Properties, and Immobilized Enzymes—cont’d Natural Polymers as Enzyme Support Properties Immobilized References

Cellulose

Longer storage capacity, better formability, and flexibility

Starch Stable (homopolysaccharide)

Fungi laccase, penicillin G acylase, glucoamylase, α amylase, tyrosinase, lipase, and β-galactosidase

Bitter gourd peroxidase

Pectin Used along with Papain (heteropolysaccharide) glycerol as plasticizer to reduce brittleness of support Pectin–chitin and pectin–calcium alginate have enhanced thermal and denaturant resistance and catalytic properties Sepharose

Al-Adhami et al. (2002), Mislovicova´, Masarova, Vikartovska, Germeiner, and Michalkova (2004), Bryjak, Aniulyte, and Liesiene (2007), Namdeo and Bajpai (2009), Labus, Turek, Liesiene, and Bryjak (2011), Huang et al. (2011), and Klein et al. (2011)

Amylase, Porous, easy adsorption, retain glucoamylase catalytic properties at extremes of pH, temperature, high salt concentration

Ceniceros et al. (2003), Go´mez et al. (2006), and Satar, Matto, and Husain (2008)

Hosseinkhani, Szittner, NematGorgani, and Meighen (2003)

Adapted from Datta, S., Christena, L. R., & Rajaram, Y. R. S. (2013). Enzyme immobilization: An overview on techniques and support materials. 3 Biotech, 3, 1–9.

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domain of chitinase A1 has high affinity toward chitin that can be exploited to retain D-hydantoinase (Chern & Chao, 2005). 2.2.1.3 Cellulose

It is the most commonly used carrier molecule for enzyme immobilization. It has lower binding capacity, inexpensive, and commercially available in fibrous and globular form. Some of the widely used enzymes which are immobilized by cellulose are listed in Table 2. Namdeo and Bajpai (2009) reported that for starch degradation, when α-amylase was immobilized to cellulose dialdehyde-coated magnetite nanoparticles, it resulted in the formation of a novel starch degrading system. Similarly, increase tolerance and feasibility of the enzyme was obtained, by immobilizing it onto ionic liquid-cellulose film activated with glutaraldehyde (Klein et al., 2011). 2.2.1.4 Collagen

The properties of collagen which make it a good support system includes proteinaceous nature, efficient water-holding capacity, and good porosity. During immobilization, a covalent bond is formed between the side chains of collagen and the enzyme, thereby strongly holding the enzyme onto the support. By using glutaraldehyde as cross-linking agent, it has been used for immobilizing tannase (Katwa et al., 1981). Chen, Song, et al. (2011) reported that when catalase was immobilized with Fe3+-collagen fibers, the activity was maintained significantly even after 26 reuses. 2.2.1.5 Carrageenans

It is a red algal linear sulfated polysaccharide. Properties of carrageenans which makes it an efficient support system includes good gelling properties, high-protein-holding capacity pseudoplastic in nature, which makes it thin under shear stress and can able to recover its viscosity once distressed. It was reported that for biodiesel production by coextrusion method using carrageenans as a support matrix, an encapsulation efficiency of 42% has been achieved (Jegannathan et al., 2010). It was found to be an inexpensive and durable carrier for lactic acid and α-galactosidase enzyme entrapment (Girigowda & Mulimani, 2006; Rao et al., 2008). 2.2.1.6 Starch

It is a natural polymer with linear amylose and branched amylopectin units with good water-holding capacity and efficient enzyme immobilizer. It was

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reported that calcium alginate–starch hybrid support was used to immobilize bitter gourd peroxidase. It was also observed that the entrapped enzyme was more stable in presence of a denaturant like urea because of internal carbohydrate moieties; on the other hand the surface-immobilized enzymes have enhanced activity (Matto & Husain, 2009). 2.2.1.7 Pectin

It is a structural polysaccharide of plants present in the primary cell wall. In plant tissues, it acts as an intercellular cementing material. It is a gelling agent with good water-holding capacity. Ceniceros et al. (2003) developed new materials for skin injury treatment by immobilizing the enzyme papain onto pectin along with 0.2–0.7% of glycerol. Pectin–chitin and pectin–calcium alginate, because of their ability to form highly stable polyelectrolyte complexes with the entrapped enzyme, the thermal and denaturant resistance and also the catalytic properties of the immobilized enzyme was significantly enhanced (Go´mez, Raml´rez, Neira-Carrillo, & Villalonga, 2006; Satar et al., 2008). 2.2.1.8 Sepharose

Sepharose is commercially available as beaded forms and is activated by CNBr. CNBr-activated Sepharose 4B used to immobilize amylase, glucoamylase which increases its porosity and easy adsorption of macromolecules. Alkyl-activated Sepharose increases the retaining catalytic property at extreme pH, high salt concentration, and high temperatures (Hosseinkhani et al., 2003). 2.2.2 Synthetic Polymers as Support Synthetic polymers are ion-exchange resins which have porous surface and are insoluble in nature. These polymers, because of their porous structure, can immobilize the enzyme very strongly. They are inert to microbial attack. They include polystyrene, polyvinyl chloride (PVC), polyacrylate, polyamide, polypropylene, diethylaminoethyl cellulose (DEAE cellulose), UV-activated polyethylene glycerol, etc. Polystyrene was first synthetic polymer used for enzyme immobilization. Some more examples of synthetic polymers used for enzyme immobilization are 1. Amberlite and DEAE cellulose used for immobilization of α-amylase (Kumari & Kayastha, 2011). 2. PVC used for immobilization of cyclodextrin glucosyltransferase, prevented thermal inactivation of the enzyme (Abdel-Naby, 1999).

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3. Synthetic polymer polyurethane microparticles was obtained by mixing polyvinyl alcohol and hexamethyl diisocyanate in the ratio of 1:3. Using polyurethane microparticles for enzyme immobilization improved enzyme loading and efficiency (Romaskevic, Vikantiene, Budriene, Ramanaviciene, & Dienys, 2010). 4. Polyethyleneglycol along with glutaraldehyde is used for immobilization of white radish peroxidase, forms a protective layer around the active center of enzyme to prevent the effect of oxidative stress (Ashraf & Husain, 2010). 5. UV-activated polyethylene glycol which has high porosity was used for waste water treatment (Xiangli, Zhe, Yinglin, & Zhengjia, 2010). 6. Glutaraldehyde-activated nylon was used for immobilizing lipase (Pahujani, Kanwar, Chauhan, & Gupta, 2008).

3. APPLICATIONS OF IMMOBILIZED ENZYMES Immobilized enzymes help in their economic reuse and also in the processing of various products. Immobilized enzymes have varied biotechnological, biomedical, and industrial applications. Furthermore, enzyme immobilization has progressed to various fields to study environmental, clinical, and industrial samples. The advancement in this area lies in the fact that they are economical, environmentally friendly, and easy to use in comparison with other parallel technologies. In the further sections, the recent developments and applications of enzyme immobilization are discussed.

3.1 Immobilized Enzymes as Biosensors The electrical, chemical, optical, or mechanical devices which can detect biological species are called biosensors. In other words, it is an analytical device which can detect and quantify specific analytes in complex samples. They are modified by biological entities like enzymes, antibodies, and oligonucleotides to increase their selectivity. The ideal biosensor is the one which has the ability to detect low concentrations of the analytes, interpret the results instantaneously and also the ability to differentiate among species according to the recognition molecules that are entrapped on its surface (Malhotra & Chaubey, 2003). Because of their exceptional properties such as high specificity, sensitivity, rapid response, low cost, compact size, user friendly, easy operation, and monitoring, make them an important tool for detection of various biological and chemical components (Amine, Mohammad, Bourais, & Palleschi, 2006). In general, biosensors have varied

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applications such as for medical diagnostics, detection of toxins in food and water, pharmaceuticals, environmental monitoring, and food and agricultural industries (Leung, Shankar, & Mutharasan, 2007). However, the development of biosensors based on enzyme immobilization has solved various challenges like enzyme stability, enzyme loss, reduced the time of enzymatic response, and offer disposable devices which can be easily used in stationary or in flow system. Diabetic patients lack insulin hormone, and so their blood glucose level remains high which has to be checked regularly to ensure that it is maintained in normal range. In the conventional method, the amount of glucose in the blood of patient is measured by the immobilized enzyme glucose oxidase and then another mediator molecule changes the color which indicates amount of glucose present. But using biosensors, instead of color, an electric current is produced which is measured by a meter to give exact concentration of glucose in blood. Biosensors (enzyme based) are also used to detect analytes such as heavy metals, organophosphorus and organochlorine pesticides, glycoalkaloids, and insecticides. It has been reported that glucose oxidase-based biosensors immobilized by electropolymerization help to detect heavy metals (Malitesta & Guascito, 2005). Similarly, urease when immobilized in both PVC and cellulose triacetate layers on the surface of pH-sensitive iridium oxide electrodes helps to detect the levels of mercury. The immobilization of polyphenol oxidase during the anodic electropolymerization of polypyrrole has been also reported. The development of biosensors for the detection of toxic pesticides is of great concern (El-Kaoutit, Bouchta, Zejli, Izaoumen, & Temsamani, 2004). Some pesticides and toxins are very harmful as they inhibit enzyme such as acetylcholinesterase A (AchE), which breaks down neurotransmitter acetylcholine. By developing biosensors containing enzyme (AchE) and its substrate (acetylcholine), the breakdown of AchE and its substrate can be detected by producing an electrical current. Immobilized enzymes which are used as biosensors for detection of compounds in food and environment are listed in Table 3.

3.2 Immobilized Enzymes in Medicine and Antibiotics Immobilized enzyme has revolutionized the medical fields, as they are tremendously used for diagnosis and treatment of various diseases, in less time, less manpower, and with high accuracy and reliability. The features of biosensors like reliability, sensitivity, accuracy, along with the unique properties

Table 3 Immobilized Enzymes Used as Biosensors for the Detection of Various Compounds in Area of Food and Environment Enzymes Inhibitors Substrate Immobilization Support References Biosensors for the determination of pesticides

Acetylcholinesterase

Paraoxon

Real water Multiwell carbon nanotubes sample

Joshi et al. (2005)

Acetylcholinesterase

Paraoxon

Orange juice

Entrapment in TCNQ-graphite

Schulze, Schmid, and Bachmann (2002)

Acetylcholinesterase

Paraoxon

Spiked

Entrapment in a PVA-SbQ

Bachmann et al. (2000)

Biosensors for the determination of heavy metals

Urease

Hg2+, Cu, Cd

Tap and Entrapment in sol–gel matrix river water

Tsai and Doongg (2005)

Urease

Hg(NO3), phenyl mercury, HgCl2, Hg2(NO3)2

Water samples

Entrapment in sol–gel film

Doong and Tsaii (2001)

Glucose oxidase

Hg2+

Spiked water

Cross-linking with GA and BSA

Mohammadi, El-Rhazi, Amine, Brett, and Brett (2002)

Glucose oxidase

Chromium (IV)

Soil sample Cross-linking with GA and Zeng, Tang, Shen, Huang, and covering with alamine membrane Niu (2004)

Biosensors for the determination of other chemical components

Butyrylcholinesterase α-Chaconine, α-solanine solanide

Potatoes

Butyrylcholinesterase α-Chaconine, α-solanine

Agriculture Cross-linking with BSA and GA vapor

Dzyadevych et al. (2003)

Butyrylcholinesterase Tomatine

Tomatoes

Dzyadevych et al. (2006)

Cross-linking with GA vapor

Cross-linking with BSA and GA

Korpan et al. (2002)

Adapted from Khan, A. A., & Alzohairy, M. A. (2010). Recent advances and applications of immobilized enzyme technologies: A review. Research Journal of Biological Sciences, 5, 565–575.

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of enzymes have led to the development of enzyme-based medicines and antibiotics (D’Orazio, 2003). In recent years, development of biosensors for the detection and measurement of metabolites is an area of intense research as it has been found that continuous metabolic monitoring provides an early indication of various body disorders and diseases. Studies showed that there is a need to replace the existing diagnostic tools such as glucose test strips, enzyme-linked immunosorbent assay, mass spectroscopy, and chromatography with faster and low cost devices that detect the early indication of any metabolic disturbance so that various disorders can be prevented at right stage (Vaddiraju, Tomazos, Burgess, Jain, & Papadimitrakopoulos, 2010). Hence the development of rapid, sensitive, and portable biosensor with immediate interpretation of the results is well adapted for this purpose. Immobilized enzymes help to identify the disease type and also the pathological conditions. Some of them are listed in Table 4. Further immobilized enzymes are also used to analyze antigens and antibodies, for examples, alkaline phosphatase, β-lactamase. Pastore and Morisi (1976) by using immobilized enzyme penicillin acylase/amidase produced 6-aminopenicillanic acid by deacylation of the side chain either in penicillin G or V. This is the most significant application of immobilized enzymes in pharmaceutical industry. These enzymes are immobilized by number of ways and can be used repeatedly. The various immobilized enzymes used for the detection of diagnostic important analytes are listed in Table 4. Table 4 Various Enzymes Used for the Detection of Diagnostic Important Analytes Enzyme Used for Diagnostic Detection Use References

Glucose oxidase

Glucose

Bostick and Hercules (1975)

Urease

Urea

Keyes and Barabino (1975) and Watson and Keyes (1976)

Cholesterol oxidase

Cholesterol

Anamika, Ajeet, and Hemant al. (2014)

Alcohol oxidase

Alcohols

Kuswandi, Irmawati, Hidayat, Jayus, and Ahmad (2014)

Penicillinase

Penicillin

Ismail and Adeloju (2010)

Aldolase

Muscle disorders

Raja, Raja, Imran, Santha, and Devasena (2011)

Adapted from Khan, A. A., & Alzohairy, M. A. (2010). Recent advances and applications of immobilized enzyme technologies: A review. Research Journal of Biological Sciences, 5, 565–575.

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3.3 Immobilized Enzymes in Food and Dairy Industry 3.3.1 Making Lactose-Free Milk In a country like India, where lactose intolerance is quite prevalent, large population can only consume lactose-hydrolyzed milk. The enzyme lactase is present in milk which hydrolyses lactose sugar to glucose and galactose. But few people lack this enzyme and cannot breakdown lactose, which is then fed by bacteria and it causes digestive problems. Lactose-free milk can be produced by immobilization techniques. Normal milk is eluted through immobilized lactase-containing column, and the milk obtained contains the products, i.e., glucose and galactose, which can easily be digested by lactose-intolerant people (Roig, BelloF, Velasco, De Celis, & Cachaza, 1987). Microbial contamination is another major problem in large-scale continuous processing of milk, so to overcome this problem, a coimmobilizate has been used. This is obtained by binding glucose oxidase on microbial cell wall using con A (Roig et al., 1987). Lactose-hydrolyzed whey may be used as whey-based beverages, feed stuff, or can be used to produce ethanol and yeast. By this way, a low cost byproduct can be converted into nutritious and quality food ingredient. 3.3.2 High-Fructose Corn Syrup The most important use of immobilized enzyme in food industry is the conversion of glucose syrups to high-fructose corn syrup (HFCS) by the enzyme glucose isomerase. The enzyme is immobilized and proceeds to the conversion of glucose into fructose makings it sweeter. In many countries, where sugar prices are high, HFCS is used as a sweetening agent (DiCosimo, McAuliffe, Poulose, & Bohlmann, 2013). 3.3.3 Fruit Juices Cell wall of the fruits contains pectins and carbohydrates which hold plant together. Immobilized pectinase, hydrolyse pectin and thus increases the amount of fruit juices with no residues of pectin (Hiteshi, Chauhan, & Gupta, 2013). Some of the immobilized enzymes used for food industry are listed in Table 5.

3.4 Immobilized Enzyme in Bioremediation/Waste Water Treatment Waste water treatment is utmost important as industries are increasing at fast pace and so their effluents in the environment. Textile, paper, leather

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Table 5 Some Immobilized Enzyme Used in Food Industry Food Substrate Immobilization Enzymes of Enzyme Support References

β-Galactosidase Lactose, whey

Bone powder

Carpio, Gonzalez, Ruales, and Batista-Viera (2000)

Pectinases

Pectin

Resin

Sarioglu, Demir, Acar, and Mutlu (2001)

Laccase

Wine, fruit juice

Silica gel

Minussi, Pastore, and Duran (2002)

Trypsin

Lactoglobulin

Cellulose

Yamamoto et al. (2005)

Lipase

Oil and grease

Ca alginate beads

Jeganathan, Bassi, and Nakhla (2006)

Adapted from Khan, A. A., & Alzohairy, M. A. (2010). Recent advances and applications of immobilized enzyme technologies: A review. Research Journal of Biological Sciences, 5, 565–575.

industry effluents are rich in dyes, which are carcinogenic even in small concentrations (Akhtar, Khan, & Husain, 2005). Bioremediation appears as a promising technique; moreover, recent studies indicate that use of enzymes for rapid degradation and removal of phenolic pollutants is an alternative to the traditional methods as well as microbial treatment that create some serious limitations (Chen & Lin, 2007). So, enzymes like peroxidase, laccase, azoreductase, etc., can play an important role to degrade these dyes. But due to harsh environmental conditions, these enzymes lose their activity, and therefore use of immobilized enzyme-based water treatment system is an ideal way. For example, HRP which is resistant to high temperature can be entrapped in calcium alginate beads (Hernandez & Fernandez-Lafuente, 2011). Immobilized laccase can also degrade a number of dyes like anthracinoid dye, lancet blue, etc. (Wu, Chou, Lupher, & Davis, 1998).

3.5 Biodiesel Production Biodiesel is a liquid fuel produced by triglycerides and esterification of alcohol in the presence of catalyst. The problem is the production of catalyst as it requires high energy. So, effort has been put for the biological production of biodiesel with the consumption of less energy. Moreover, production of biodiesel by immobilization of lipase further reduces the cost as it can be used repeatedly and has more stability (DiCosimo et al., 2013; Khan & Alzohairy, 2010).

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3.6 Immobilized Enzymes in Biotech Cleaning Immobilized enzymes are nowadays used in detergent industries for the removal of stains as they have good cleaning properties as compared to synthetic detergents (Hasan, Shah, Javed, & Hameed, 2010). These enzymebased detergents are pollution free as they do not effect environment and used in less amount as compared to synthetic detergents; further, they also work well in low temperature. These enzymes are used as detergents after immobilizing it on suitable substrates to overcome few problems like stability, cost, repeated use, etc.

3.7 Carbon Capture The most alarming problem of environment is significant high levels of carbon dioxide in air. Majority of CO2 comes from industrial power plants. Intense research is going on to convert or capture carbon dioxide coming from power plant into useful material or store in underground (Rubin, Mantripragada, Marks, Versteeg, & Kitchin, 2012; Yu, Curcic, Gabriel, & Tsang, 2008). The enzyme carbonic anhydrase is one of the fastest enzymes that catalyzes the hydration of CO2 bicarbonate ðHCO3
Þ and protons (H+), with hydration turn over number around 106 s
1. The catalytic efficiency of this enzyme is dependent on the proton shuttle which is nearly the diffusion limit in buffers with pKa >8. Hence this is the ideal enzyme for CO2 capture because of its fastest kinetics as well as its potential to identify enzymes that catalyzes the reaction under varied pH and temperature conditions (Yu et al., 2008). Therefore immobilization of this enzyme in power plant chimney can directly extract the CO2 from the smoke and convert it into bicarbonate. Hence, the smoke that leaves the plant will have only bicarbonate, which can be later converted back into CO2 and can be stored. However, the disadvantage of using carbonic anhydrate is that it denatures at high temperatures. So, intense research is going on this field in order to modify this enzyme to resist high temperatures of the power plant (DiCosimo et al., 2013).

4. CONCLUSIONS In recent years, significant progress in design of enzyme immobilization, support matrix with different pore size, and surface modifications are developed. Designing ideal support material by modifying specific structural

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features required for a target enzyme is now possible by new simulations. Moreover, as the structure of enzyme and the mechanism of action is known, controlled immobilization methods can be developed. Furthermore, identification and characterization of essential metabolites and their screening for curing diseases have received significant attention. This is mainly because of development of economical and disposable array biosensors, biochips, and bioreactors. It is our view that the future holds significant promise with increased usage of immobilized enzymes in pharmacological, clinical, food, biotechnological, and other industrial fields.

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INDEX Note: Page numbers followed by “f ” indicate figures, and “t” indicate tables.

A Acrylamide, 2–3, 12–13 Actinobacterial enzymes, 68 aminoacylase, 84 biocatalysts, 71, 86 chitinase, 85–86 extremophilic enzymes, 71–73, 72t lignocellulolytic enzymes, 69, 77–84 lipase, 86 oxidative enzymes, 69, 73–77, 78–79t penicillin amidase, 86 protease, 84–85 Actinomycetes, 68–71 Activated carbon, 190–191 Adsorption/carrier-binding method, 181–183 Aeromonus hydrophila, 49 Affinity binding, enzyme immobilization, 184–185 Agar, 107 Alginate block structures, 138 in commercial food industries, 138 enzyme immobilization, 191 mannuronate chemical structures, 139f oligosaccharides, 138 in pharmaceutical industries, 138 polymers, 138 Alginate lyase bifunctional alginate lyase, 151–153 bioactivity, 153–154 enzymological studies, 140–142 growth-promoting effects, 155–157 guluronate lyase, 148–151 immune systems activation, 153–155 mannuronate lyase, 142–148 marine molluscs, 140–142 plaque formation assay, 140–142, 141f spectrophotometric assay, 142f viscosity of alginate solution, 141f Alginate oligomer mixture (AOM), 154–155 Alkaline protease, 104

L-Amino acid oxidases (L-AAO), 76 Aminoacylase, 84 Amylase, 107 advantages, 162–163 characterization, 168–169 dextrinizing activity, 170–171 dinitrosalicylic acid method, 170 enzyme activity determination, 169–171 Indian pharmacopeia method, 171 industrial applications, 171–173 molecular biology, 169 Nelson–Somogyi method, 170 production, 164–167 purification, 167–168 starch suspension, 171 types, 163–164 using iodine, 170 α-Amylase, 163 production, 165–166 β-Amylase, 163–164 production, 166–167 Amylograph/Farinograph test, 171 Antibiotics, enzyme immobilization, 197–199 Aplanochytrium sp., 101 Aplysia kurodai, 142–144 Archaeoglobus fulgidus, 46 Arthrospira platensis, 8–11 L-Asparaginase, 2, 57–59 application, 2 bacterial type, 5t enzymatic hydrolysis, 13 from Erwinia chrysanthemi, 2 from Escherichia coli, 2 in food, 12–13 history, 2–3 marine environment, 2 challenges, 16–20 microorganisms, 15–16, 17–19t ocean basin, 14–15, 15f potential of, 14–15 in medicine, 11–12 213

214 L-Asparaginase (Continued ) multiple alignment sequences, 6f occurrence and distribution, 3–4, 4t optimization, 9–10t production, 8–11, 50–53 structure and properties, 4–7, 6–7f Asparaginase I (AnsI), 4 Asparaginase II (AnsII), 4 Asparagine, 11–12 Aspergillus A. falvus var. columnaris, 167–168 A. niger, 13, 165 Aureobasidium pullulans, 36–37, 54, 167 Autonomous underwater vehicles (AUVs), 19

B Bacillus sp., 165–166 B. aryabhattai, 16 B. brevis, 8–11 B. mojavensis, 57–59 B. subtilis, 13, 108–109 Bacterial enzymes, 104–108, 105–106t Bakery industry, amylases in, 172 Barophilic extremozymes, 49–50 Bifunctional alginate lyase, 151–153 characterization, 152–153 purification, 152 Biocatalysis process, 180–181, 183 Biodiesel production, 201 Bioremediation, 200–201 Biosensors, 196–197, 198t Biotech cleaning, 202

C Cancer therapy, 2, 11–12 Carbon capture, enzyme immobilization, 202 Carrageenans, 194 Catechol 1,2-dioxygenase, 74–75 Cellulases, 69, 80 classification, 30–31 global enzyme market, 30 industrial uses of, 29f marine microbes, 28–30, 32–33t bacteria, 31–35 fungi, 35–36 source, 31 yeast, 36–37

Index

SSF-derived, 35–36 Cellulose, 108, 194 Ceramics, 190 Chaetomium globosum, 109 Charcoal, 190 Chitin, 107 Chitinase, 85–86 Chitinoglastic bacteria, 107 Chitosan, 191–194 Chlamydomonas reinhardtii, 138–140, 156–157 Cholesterol oxidases (CO), 73–74 Clostridium C. acetobutylicum, 168 C. thermohydrosulfuricum, 169 Cold adaptation, 118, 120–122, 124 Cold-adaptive enzymes, 46–47 Collagen, enzyme immobilization, 194 Corn syrup, high-fructose, 200 Covalent binding method, 183–184 Creatine kinase (CK), 124–128, 125f, 127f Cyclic AMP (cAMP), 11

D Dairy industry, enzyme immobilization, 200 Deep-sea hydrothermal vents, 46 Degree of polymerization (DP), 145–148 Dehydroepiandrosterone sulfate (DHEAS), 73–74 Desulfovibrio desulfuricans ATCC27774, 109 Dinitrosalicylic acid (DNS) method, 170 DrLDH, 122–123

E ED-AOM, 156–157 Entrapment method, 185–186 Enzymatically digested alginate oligomers, 153–157 Enzymes bacterial, 104–108, 105–106t cold-adaptive, 46–47 fungal, 101–104 immobilization (see Immobilization of enzyme) marine microbes, 28 salt-active, 48–49 volume, 120

215

Index

Erwinia chrysanthemi, 2, 5 Escherichia coli, 2, 5 E. coli L-asparaginase (EcIII), 7 extremozymes, 46–47, 50–55 Extremophiles, 45, 57 from actinobacteria, 71–73, 72t Extremozymes, 45, 71, 72t barophilic, 49–50 halophilic solvent active, 48–49 industrial applications, 57–59 large-scale production, 56–57 marine actinobacteria, 50–53 microbial enzymes, application of, 53–55 organic solvent active, 48–49 pH stability, 48 psychrophilic, 46–47 structural elucidation, 55–56 thermophilic, 45–46

F Falling number (FN) method, 171 Fatty acids desaturase, 129–131 Fish, 118, 124f creatine kinase, 124–128, 125f, 127f fatty acids desaturase, 129–131 lactate dehydrogenase, 119–124, 119f, 121f, 124f Flammeovirga, 148–149 Food industry L-asparaginase, 12–13 enzyme immobilization, 200, 201t Food safety, 12–13 Free energy, 120 Fructose, 171–172 Fruit juices, 200 Fungal enzymes, 101–104

G Gillichthys seta, 120–122 Glass, enzyme immobilization, 190 Glucoamylase, 164, 167 Glucose, 171–172 L-Glutamate oxidases, 76–77 L-Glutaminase, 57–59 Green enzymes, 73 6 M Guanidine hydrochloride (GHCl), 150–151

Guluronate lyase amino acid compositions, 151 characterization, 150–151 molecular weight, 151 purification, 148–150

H Halobacterium salinarum, 48–49, 165–166 Halophilic extremozymes, 48–49 Hemicellulases, 69, 80–82 Hemicelluloses, 77–79 Homeoviscous adaptation hypothesis, 129 Horseradish peroxidase (HRP), 74 Hydrothermal vents, deep-sea, 46

I Immobilization of enzyme adsorption/carrier-binding method, 181–183 affinity binding, 184–185 Bacillus circulans, 191–194 biodiesel production, 201 bioremediation/waste water treatment, 200–201 biosensors, 196–197, 198t biotech cleaning, 202 carbon capture, 202 covalent binding method, 183–184 in dairy industry, 200 definition, 180–181 entrapment method, 185–186 in food industry, 200, 201t fruit juices, 200 high-fructose corn syrup, 200 ionic binding method, 186 lactose-free milk, 200 medicine and antibiotics, 197–199 metal-linked method, 186 methods, 180–181, 182f nanodiametric supports, 184 support material, 187 activated carbon, 190–191 alginate, 191 carrageenans, 194 cellulose, 194 ceramics, 190 charcoal, 190 chitosan, 191–194

216 Immobilization of enzyme (Continued ) collagen, 194 glass, 190 inorganic material, 188–191, 189t natural polymers, 191–195 organic supports, 191–196, 192–193t pectin, 195 sepharose, 195 silica, 188–190 starch, 194–195 synthetic polymers, 195–196 Immune systems activation, alginate lyase, 153–155 Indian pharmacopeia method, 171 Iodine, 170 Ionic binding method, 186

L Lactate dehydrogenase (LDH), 119–124, 119f, 121f, 124f Lactose-free milk, 200 Lignocellulolytic enzymes from actinobacteria, 69, 77–84 from marine fungi, 102–103t Lignolytic enzymes, 82–84 Lipases, 86, 106–107 biodiesel, 201 Candida antarctica, 190 Candida rugos, 183 cold-active, 47 marine microbes, 57–59 Yarrowia lipolytica, 183 Lipid composition, 129 Liquefaction, 171–173 Lupinus luteus (L1A), 7

M Mangrove ecosystem, 99–101 bacterial enzymes, 104–108, 105–106t fungal enzymes, 101–104 Mannuronate lyase characterization, 145–148 circular dichroic spectrum, 147f molecular weight, 145–148, 146t purification, 142–144 substrate recognition site, 147f substrate-specific actions, 143f Marine actinobacteria, 50–53 Marine fungi, 102–103t

Index

Marine microorganisms, 28–30, 100–101 cellulases, 32–33t bacteria, 31–35 fungi, 35–36 source, 31 yeast, 36–37 diversity, 28 environment diversity, 162 enzymes, 28, 30, 44–45 esterases and lipases, 57–59 features, 44 thermotolerant alkaliphilic proteases, 48 Marine polysaccharases, 28, 29f Marinobacter sp., 50–53 Medicine L-asparaginase, 11–12 enzyme immobilization, 197–199 Metal-linked method, 186 Methanococcus jannaschii, 49–50 Microbial contamination, 200 Microbial enzymes application, 53–55 from marine fungi, 102–103t Microbispora siamensis, 80–81 Micrococcus agilis, 84 Mucor sp., 165–166 Mycobacterium smegmatis, 84

N Nannochloropsis oculata, 155–156 Nelson–Somogyi method, 170 Nocardiopsis N. aegyptia, 50–53 N. prasina, 85–86

O Ocean basin, 14–15, 15f Ocean pressure, 15 Oxidative enzymes, from actinobacteria, 69, 73–77, 78–79t

P Pectin, enzyme immobilization, 195 Penicillin amidase, 86 Peroxidases, 74 pH stability, extremozymes, 48 Pichia pastoris, 79–80 Plackett–Burman design, 56–57

217

Index

Plant cells, growth-promoting effect, 155–157 Polyhydroxybutyrate depolymerase, 50–53 Poly(α-L-guluronate) lyase (PG lyase), 140–142 Poly(β-D-mannuronate) lyase (PM lyase), 140–142 Polysaccharide-degrading enzymes, marine microbes, 28 Polysaccharide lyase 7 (PL7), 148–149 Preussia minim, 167–168 Protease, 84–85, 104 Pseudoalteromonas P. arctica, 47 P. atlantica, 109 Pseudoalteromonas sp. strain No. 272, 152 Pseudomonas fluorescens, 148 Psychrophilic extremozymes, 46–47 Putrescine oxidase (PuOs), 76 Pyrococcus furiosus, 46, 49, 53–54, 168–169

R RAW264.7 cells, 153–155 Red seaweeds, 107–108 Remotely operated vehicles (ROVs), 19 Rhizobacterium etli, 4 Rhizophora apiculata, 101 Rhodococcus erythropolis, 86

S Saccharification, 34–36 Saccharopolyspora sp. A9, 165–166 Saturated fatty acids (SFAs) desaturation, 129–130 Schizochytrium aggregatum, 101 Sclerotium rolfsii, 85–86 Sepharose, 195 Silica, enzyme immobilization, 188–190 Small laccase (SLAC), 83–84 Solid-state fermentations (SSF), 8, 35–36, 164–166 Starch enzymatic hydrolysis, 171–172 enzyme immobilization, 194–195 gelatinization, 171–172 suspension, 171 Stearoyl-CoA desaturase (SCD), 129–131, 129f, 131f Streptococcus bovis 148, 168–169

Streptomyces sp., 165–166 S. albidoflavus, 71–72 S. albus, 81–82 S. aureofaciens, 80–81 S. castaneoglobisporus, 75 S. coelicolor, 80–81 S. flavogriseus, 81–82 S. fungicidicus, 50–53 S. hygroscopicus, 81–82, 85–86 S. lividans, 79–80 S. pseudogriseolus, 81–82 S. reticuli, 79–80 S. thermoviolaceus OPC-520, 85–86 Submerged fermentations (SmF), 8, 164–166 Super-high frequency (SHF) radiation, 70–71

T Temperature, 118–119 Thermoactinomyces, 81–82 Thermobifida fusca, 71–72 Thermococcus T. aggregans, 46 T. hydrothermalis, 46 T. kodakaraensi gene (TK1656), 11 Thermophilic extremozymes, 45–46 Thermotoga maritime, 109 Thermus aquaticus, 46, 53–54, 109 Threonine residue in the position of 12 (Thr12), 5 Topoisomerases, 109 Triacylglycerol hydrolases. See Lipases TtLDH, 122–123 Twin-arginine translocation (TAT) systems, 80 Tyrosinase, 75

W Wastes or underutilized materials (WUM), 8 Waste water treatment, 200–201

X Xylanase, 108

Y Yarrowia lipolytica, 54, 183

Z Zunongwangia profunda, 46–47