Marine Enzymes Biotechnology: Production and Industrial Applications, Part III - Application of Marine Enzymes [1st Edition] 9780128097281, 9780128095874

Table of contents :
Content:
Series PagePage ii
CopyrightPage iv
ContributorsPages ix-x
PrefacePages xi-xiiS.-K. Kim, F. Toldrá
Chapter One - Marine Enzymes in Cancer: A New ParadigmPages 1-14R.H. Prabhu, K.S. Bhise, V.B. Patravale
Chapter Two - Bacillus Probiotic Enzymes: External Auxiliary Apparatus to Avoid Digestive Deficiencies, Water Pollution, Diseases, and Economic Problems in Marine Cultivated AnimalsPages 15-35Jorge Olmos Soto
Chapter Three - Characterization and Applications of Marine Microbial Enzymes in Biotechnology and Probiotics for Animal HealthPages 37-74T.H. Nguyen, V.D. Nguyen
Chapter Four - Biotechnological Applications of Marine Enzymes From Algae, Bacteria, Fungi, and SpongesPages 75-106S. Parte, V.L. Sirisha, J.S. D’Souza
Chapter Five - Biomedical Applications of Enzymes From Marine ActinobacteriaPages 107-123K. Kamala, P. Sivaperumal
Chapter Six - Production of Enzymes From Agricultural Wastes and Their Potential Industrial ApplicationsPages 125-148S. Bharathiraja, J. Suriya, M. Krishnan, P. Manivasagan, S.-K. Kim
Chapter Seven - Marine Enzymes: Production and Applications for Human HealthPages 149-163T. Eswara Rao, M. Imchen, R. Kumavath
Chapter Eight - Bioremediation of Industrial Waste Through Enzyme Producing Marine MicroorganismsPages 165-179P. Sivaperumal, K. Kamala, R. Rajaram
Chapter Nine - Marine Enzymes and Microorganisms for Bioethanol ProductionPages 181-197M.R. Swain, V. Natarajan, C. Krishnan
Chapter Ten - Enzymes in Fermented FishPages 199-216Giyatmi, H.E. Irianto

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 2017 Copyright © 2017 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: 978-0-12-809587-4 ISSN: 1043-4526 For information on all Academic Press publications visit our website at https://www.elsevier.com/books-and-journals

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

CONTRIBUTORS S. Bharathiraja CAS in Marine Biology, Annamalai University, Porto Novo, India K.S. Bhise Institute of Chemical Technology, Mumbai, Maharashtra, India J.S. D’Souza UM-DAE Centre for Excellence in Basic Sciences, Mumbai, India T. Eswara Rao Central University of Kerala, Padannakkad, Kerala, India Giyatmi Jakarta Sahid University, Jakarta Selatan, DKI Jakarta Province, Indonesia M. Imchen Central University of Kerala, Padannakkad, Kerala, India H.E. Irianto Center for Fisheries Research and Development, Jakarta Utara, DKI Jakarta Province, Indonesia K. Kamala Center for Environmental Nuclear Research, Directorate of Research, SRM University, Kattankulathur, India S.-K. Kim Marine Bioprocess Research Center; Specialized Graduate School Science & Technology Convergence, Pukyong National University, Busan, Republic of Korea C. Krishnan Indian Institute of Technology Madras, Chennai, India M. Krishnan School of Environmental Sciences, Bharathidasan University, Tiruchirappalli, India R. Kumavath Central University of Kerala, Padannakkad, Kerala, India P. Manivasagan Marine Bioprocess Research Center, Pukyong National University, Busan, Republic of Korea V. Natarajan Indian Institute of Technology Madras, Chennai, India T.H. Nguyen Faculty of Food Technology, Nha Trang University, Nha Trang, Vietnam V.D. Nguyen Institute of Biotechnology and Environment, Nha Trang University, Nha Trang, Vietnam ix

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Contributors

Jorge Olmos Soto Molecular Microbiology Laboratory, Centro de Investigacio´n Cientı´fica y de Educacio´n Superior de Ensenada (CICESE), Ensenada, Baja California, Mexico S. Parte UM-DAE Centre for Excellence in Basic Sciences, Mumbai, India V.B. Patravale Institute of Chemical Technology, Mumbai, Maharashtra, India R.H. Prabhu Institute of Chemical Technology, Mumbai, Maharashtra, India R. Rajaram Bharathidasan University, Tiruchirappalli, India V.L. Sirisha UM-DAE Centre for Excellence in Basic Sciences, Mumbai, India P. Sivaperumal Center for Environmental Nuclear Research, Directorate of Research, SRM University, Kattankulathur, India J. Suriya School of Environmental Sciences, Bharathidasan University, Tiruchirappalli, India M.R. Swain Indian Institute of Technology Madras, Chennai, India

PREFACE In the last decades, progress in the knowledge of marine enzymes has advanced exponentially. The growing interest on marine enzymes is related to their relevant 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, and easy adaptation to cold conditions. 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 publishes three consecutive volumes focused on the topic Marine Enzymes Biotechnology: Production and Industrial Application. Parts I and II were published in 2016 and the third part is published in 2017. Volume 78 corresponded to Part I that was mainly dealing with the production of enzymes from marine sources, volume 79 corresponded to Part II dealing with the marine organisms producing enzymes, and this volume 80 corresponds to Part III dealing with the applications of marine enzymes. This volume brings 10 chapters reporting the applications of enzymes produced by marine organisms. So, this includes the potential use of enzymes derived from marine sources as therapeutic agents for cancer therapy or other potential biological applications relevant to human health as well as the biomedical applications of enzymes from marine actinobacteria, the biotechnological applications of marine enzymes from algae, bacteria, fungi, and sponges, the use of enzymes from Bacillus subtilis to facilitate the nutrients assimilation from unconventional and economic plant resources in aquaculture marine animals, the use of marine probiotic enzymes to improve host digestion and cleave molecular signals involved in quorum sensing in pathogens to control disease in aquaculture, the economic production of actinobacterial enzymes from agricultural wastes as a better alternative for utilization of biomass, the action of marine enzymes for bioremediation of industrial wastes and for the development of efficient processes for bioethanol production, and the use of marine enzymes in

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fermented fish products. This volume presents the combined effort of 30 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. S.-K. KIM AND F. TOLDRA´ Guest Editors

CHAPTER ONE

Marine Enzymes in Cancer: A New Paradigm R.H. Prabhu, K.S. Bhise, V.B. Patravale1 Institute of Chemical Technology, Mumbai, Maharashtra, India 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Marine Enzymes as Anticancer Agents 2.1 L-Arginase 2.2 L-Arginine Deiminase 2.3 L-Asparaginase 2.4 Glutaminase-Free L-Asparaginase 2.5 L-Glutaminase 2.6 Protease 2.7 Lysozyme 2.8 Acetylcholinesterase 2.9 Laccase 2.10 Fucoidanase 3. Conclusion Acknowledgments References

2 3 5 6 6 7 8 9 9 10 11 11 12 12 12

Abstract Over the last decades, the vast chemical and biodiversity of marine environment has been identified as an important source of new anticancer drugs. The evolution of marine life is a result of competition among microorganisms for space and nutrients in the marine environment, which drives marine microorganisms to generate diverse enzyme systems with unique properties to adapt to harsh conditions of ocean. Therefore, marine-derived sources offer novel enzymes endowed with extraordinary properties. Recent advances in cancer therapy have facilitated enzyme therapy as a promising tool. But, the available information on the use of enzymes derived from marine sources as therapeutic agents for cancer therapy is scanty. The potential utility of marine enzymes in cancer therapy will be discussed in this chapter.

Advances in Food and Nutrition Research, Volume 80 ISSN 1043-4526 http://dx.doi.org/10.1016/bs.afnr.2016.10.001

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2017 Elsevier Inc. All rights reserved.

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1. INTRODUCTION Cancer is characterized by uncontrolled growth and spread of abnormal cells. Malfunction of genes that control cell growth and division is the key reason for manifestation of all types of cancer. Most cancer treatment protocol relies majorly on chemotherapy; however, chemotherapeutic drugs lack tumor selectivity affecting normal cells and also causing multidrug resistance. These setbacks result in serious side effects, immunity suppression, and poor treatment outcomes. Hence the search for novel drugs is still a priority goal for cancer therapy. Natural products have been a major source of new drug candidates since ages (Demain & Sanchez, 2009; Gupta, Pandotra, Sharma, Kushwaha, & Gupta, 2013; Prabhu, Patravale, & Joshi, 2015). Nature has been an inexhaustible source with a remarkable chemical diversity of organisms living on earth. Oceans occupy almost 80% of the world’s biota, and it is believed that life originated from the ocean. Marine ecosystem comprises of microorganisms, algae, fungi, seaweeds, sponges, fishes, and invertebrates. The demand to explore therapeutic agents in the marine environment has been outstanding in the past few years, despite the abundance of plant and animal diversity in terrestrial life ( Jeon, Samarakoon, & Elvitigala, 2015; Suleria, Osborne, Masci, & Gobe, 2015). Marine ecosystem represents a wide and extraordinary resource of biologically active metabolites because of the chemical and biological diversity of the marine environment (Wang, Liu, Su, Sun, & Wang, 2009). The fact that marine environment, predominantly sea water, is saline in nature and chemically closer to human blood plasma anticipates the possibility to find enzymes that are compatible and nontoxic to humans (Unissa, Sudhakar, & Reddy, 2016). Most of sponges, seaweeds, crustaceans, fish species, and their associated microorganisms have evolved chemical means in order to protect themselves against predation and survive in the harsh and complex marine environment. Such chemical and biological adaptation enables production of bioactive substances with beneficial effects on human health, including potential anticancer agents (Kerr & Kerr, 1999). Recently improvements in deep-sea collection and aquaculture technology are responsible for the growing interest of the enormous biodiversity present in the marine world which has led to shift of focus toward oceans as being potential source of new anticancer candidates. This has resulted in marinederived compounds entering preclinical and early clinical development phases (Mayer et al., 2010; Schwartsmann, da Rocha, Mattei, & Lopes, 2003).

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Marine-derived compounds are more bioactive in terms of cytotoxicity than those of terrestrial origin. The fact that many marine compounds have already entered into the market proves that marine natural products have their stronghold in the area of anticancer chemotherapy. The global marine pharmaceutical pipeline consists of four approved anticancer pharmaceuticals in clinical use which includes Cytarabine (Cytosar-U®; leukemia), Trabectedin (Yondelis®; soft tissue carcinoma), Eribulin (Halaven®; breast cancer), and Brentuximab vedotin (Adcetris®; Hodgkin’s lymphoma) (Crawford et al., 2016; Martins, Vieira, Gaspar, & Santos, 2014; Newman & Cragg, 2016). Enzymatic therapy is one of the promising modalities for treatment of cancer, since several enzymes are known to interfere with the growth and proliferation of metastatic cells through various mechanisms. Cancer cells are auxotrophic in nature as they exhibit enhanced requirements for essential nutrients, thereby rendering them as susceptible targets. Thus the use of enzymes to deprive cancer of essential nutrients offers a selective approach for treatment of tumor malignancies (Saxena, 2015). Marinederived enzyme has unique activity that differs from enzyme produced by other sources because marine species have evolved mechanisms to survive in an extremely hostile environment in terms of light, salinity, and pressure as compared with land (Crawford et al., 2016). Table 1 provides an overview of anticancer enzymes derived from marine sources. This chapter briefs about marine-derived enzymes with respect to its anticancer efficacy.

2. MARINE ENZYMES AS ANTICANCER AGENTS The criterion for an enzyme to be ideally suited for an anticancer therapeutic agent is that it should be produced from nonpathogenic and easily available organism strain; provide higher yield when grown on simple nutrient media; be nontoxic, stable at physiological pH and temperature; and exhibit broad range of activity against cancer cell lines (Unissa et al., 2016). Therapeutic enzymes obtained from terrestrial bacterial sources that are currently used for the treatment of cancer are known to cause several side effects, and hence, this necessitates the use of alternative enzyme drug that is nonimmunogenic, nontoxic, and compatible to human blood (Senthil, Selvam, & Singaravel, 2012). The major mechanisms by which several marine enzymes, as enlisted in Fig. 1, have been found to exert its anticancer activity by means of nutrient depletion, antiangiogenesis, or induction of

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Table 1 Marine Source of Enzymes with Anticancer Activity Enzyme Marine Source References L-Arginase

L-Arginine deiminase

L-Asparaginase

Glutaminase-free L-asparaginase

L-Glutaminase

Penicillium chrysogenum

El-Sayed, Shindia, Diab, and Rady (2014)

Idiomarina sediminum

Unissa, Sudhakar, and Reddy (2015)

Vibrio alginolyticus 1374

Unissa et al. (2016)

Aspergillus fumigatus

El-Sayed, Hassan, and Nada (2015)

Marine actinomycetes (Streptomyces species)

Dhevagi and Poorani (2006) and Basha, Rekha, Komala, and Ruby (2009)

Streptomyces acrimycini NGP

Selvam and Vishnupriya (2013)

Streptomycete species WS3/1

Kumari, Sankar, and Prabhakar (2011)

Bacillus tequilensis PV9W

Shakambari et al. (2016)

Enterobacter cloacae

Husain, Sharma, Kumar, and Malik (2016)

Penicillium brevicompactum Elshafei et al. (2014) NRC 829 Alcaligenes faecalis KLU102

Pandian, Deepak, Sivasubramaniam, Nellaiah, and Sundar (2014)

Vibrio azureus JK-79 (JQ820323)

Kiruthika and Saraswathy (2013)

Protease

Bacillus species

Vijayasurya, Tintu, and Manjusha (2014)

Lysozyme

Marine bacilli

Ye, Wang, Chen, Guo, and Sun (2008)

Acetylcholinesterase Spirastrella pachyspira, Halichondria glabrata, and Cliona lobata

Kumar and Gopalkrishnan (2014)

Laccase

Cerrena unicolor

Matuszewska et al. (2016)

Fucoidanase

Streptomyces species

Manivasagan and Oh (2015)

Pseudoalteromonas citrea KMM 3296 and KMM 3298

Bakunina et al. (2002)

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Marine Enzymes in Cancer: A New Paradigm

Nutrient depletion L-Arginase L-Arginine

deiminase

L-Asparaginase L-Glutaminase Protease

Anticancer mechanism of marine enzymes

Antiangiogenesis Lysozyme Acetylcholinesterase

Apoptosis Laccase Fucoidanase

Fig. 1 Mechanism of anticancer activity of marine enzymes.

apoptosis (Ye et al., 2008). The details of marine-derived enzymes which have been explored for its anticancer potency will be discussed further.

2.1 L-Arginase L-Arginase is a powerful anticancer, L-arginine-depleting enzyme. It is active against argininosuccinate synthase expressing tumors by hydrolyzing L-arginine to L-ornithine and urea (Philip, Campbell, & Wheatley, 2003). L-Arginase obtained from submerged fungi cultures of thermotolerant Penicillium chrysogenum exhibited a plausible thermal stability at 40°C and threefold higher affinity to L-arginine (Km 4.8 mM) than PEG-arginase (PEG-arg) (Km 15.2 mM). From the cytotoxicity assay, it was found that the arginase enzyme has shown higher activity against hepatic cellular carcinoma (HEPG-2) (IC50 0.136 U/mL) and human lung carcinoma (A549) (IC50 0.165 U/mL) tumors, by about twofold in comparison to PEG-arginase. This enhanced anticancer activity of arginase confirms its arginine auxotrophic identity. The biological half-life of free and PEG-arginase when assessed as holoenzymes was found to be 16.4 and 20.4 h, respectively, while on addition of cofactor Mn+, their activities were found to increase by 1.8to 2-folds. The free enzyme was found to slightly induce the titer of immunoglobulins (IgGs) by 10–15% after 28 days of injection in rabbit model in comparison to the PEG-arg that had shown relatively unchanged IgG titers (El-Sayed et al., 2014).

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2.2 L-Arginine Deiminase L-Arginine deiminase (ADI) is a therapeutic enzyme that acts on various arginine auxotrophic cancer cells that do not express ornithine carbamoyl transferase and argininosuccinate synthetase. In most cases, cancer cells require continuous supply of L-arginine for their growth and survival. They utilize this amino acid as a source of nitrogen for the synthesis of their cell components. Demand for L-arginine by cancer cells is higher than normal cells, and its depletion can starve cancer cells to death (Bowles et al., 2008). ADI has been effectively applied in therapy of cancer as nutrient depleter. Unissa et al. (2016) evaluated the antiproliferative activity of the purified ADI obtained from novel marine bacterial isolate Vibrio alginolyticus 1374 against various human cancer cell lines. The enzyme exhibited IC50 values of about 5.21, 6.3, 8, and 3.13 U/mL when tested on A375-C6 (human melanoma), MCF-7 (human breast cancer), HCT-113 (human colon carcinoma), and Jurkat, clone E6-1 (human T lymphoblast) cell lines, respectively. The antiproliferative activity of enzyme on Jurkat, clone E6-1 cell lines was higher than other cell lines tested (Unissa et al., 2016). To increase thermostability of the enzyme, ADI sourced from thermophilic Aspergillus fumigatus KJ434941 was PEGylated and covalently immobilized on dextran (DEX). PEG-ADI and DEX-ADI displayed thermostability 2-fold more at 50°C and 1.7-fold more at 70°C than the free enzyme. The free enzyme and PEG-ADI exhibited similar anticancer efficacy but lower than DEX-ADI when tested against HCT, HEP-G2, and MCF7. The IC50 values of free ADI were 22.0, 16.6, and 13.9 U/mL, while for DEX-ADI were 3.98, 5.18, and 4.43 U/mL for HCT, MCF7, and HEPG-2, respectively. Thus the covalent modification by dextran was found to increase the anticancer activity of the enzyme by five-, three-, and threefold against HCT, MCF7, and HEPG-2, respectively. The pharmacokinetic evaluation revealed that the in vivo half-life of free ADI, PEG-ADI, and Dex-ADI was 29.7, 91.1, and 59.6 h, respectively. The biochemical and hematological parameters were not affected by dosing of free or modified ADI and no signs of antigenicity were detected. This thermostable and nonantigenic ADI proves to be potential candidate for preclinical anticancer research (El-Sayed et al., 2015).

2.3 L-Asparaginase The auxotrophic type of cancer cells requires L-asparagine for its essential cellular processes. L-Asparaginase enables the depletion of existing

Marine Enzymes in Cancer: A New Paradigm

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L-asparagine by converting L-asparagine to L-aspartic acid and ammonia, and

thus forces the tumor cells to undergo apoptosis (Savitri & Azmi, 2003). isolated from Streptomyces species obtained from marine actinomycetes was explored for antilymphoblastic leukemia activity. The enzyme exhibited maximum activity between pH 8 and 9 at 60°C. The L-asparaginase inhibition effects on the leukemic cell types were evaluated and compared with Topotecan (clinical drug for leukemia therapy) and PD18435214 (cytotoxic molecule). It showed cytostatic effect on JURKAT cells (acute T cell leukemia) and K562 cells (chronic myelogenous leukemia) at 24 h; the growth inhibition (GI50) evidenced was similar to the cytostatic effects of Topotecan and higher than PD184352. At 48 h, L-asparaginase showed cytotoxic effects on the leukemic cell lines comparable to the growth inhibition produced by Topotecan. L-Asparaginase-induced cell cycle arrest and apoptosis of leukemia cells were considered as the possible anticancer mechanisms of this enzyme (Dhevagi & Poorani, 2006). Marine actinomycete Streptomyces acrimycini NGP was utilized as a source of L-asparaginase, and the antioxidant and cytotoxic potential of the enzyme were assessed. The antioxidant study of this enzyme revealed good antioxidant activity against H2O2 and DPPH with an IC50 value of 78.8 and 63.3 μg/mL, respectively. In vitro anticancer activity performed against gastric stomach cancer (AGS) cell line showed that L-asparaginase inhibits the AGS cells with an IC50 of 49.11 μg/mL. The antioxidant potential of this enzyme also plays an influential role in its anticancer efficiency (Selvam & Vishnupriya, 2013). L-Asparaginase

2.4 Glutaminase-Free L-Asparaginase The glutaminase-free property of L-asparaginase is essential to avoid the reduction of L-glutamine from the circulatory system, which results in severe side effects in patients like seizures, hyperglycemia, leucopenia, and acute pancreatitis (Manna, Sinha, Sadhukhan, & Chakrabarty, 1995). A marine bacterial isolate Bacillus tequilensis PV9W was evaluated as a potential source of glutaminase-free L-asparaginase. The purified enzyme had an apparent Km of 0.045  0.013 mM and Vmax of 7.465  0.372 μmol/mL/min values. The maximum activity of L-asparaginase was achieved at a pH 8.5 and 35°C. This enzyme exhibited effective cytotoxicity with low IC50 value of 0.036  0.009 IU/mL against human cervical cancer cell lines (HeLa). Further mechanistic investigation of L-asparaginase-treated HeLa cells by flow cytometry revealed G2 arrest in cervical cancer cells by modulating

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p53 expression. Arrest of cells in G2 phase was attributed to deprivation of L-asparagine. The p53 expression in L-asparaginase-treated HeLa cells was found to be higher than untreated control cells by blocking the transition of the cells into mitosis phase. This enzyme was found to be nonhemolytic and had higher in vitro trypsin half-life (42.59 min), thereby paving its way as a potential anticancer oncolytic enzyme (Shakambari et al., 2016). The glutaminase-free asparaginase produced from Enterobacter cloacae was assessed for cytotoxicity against a panel of human cancer cell lines, viz., promyelocytic leukemia cell line (HL60), T lymphoblast acute lymphoblastic leukemia cell line (MOLT-4), and breast cancer cell lines (MDA-MB231, T47D). Highest cytotoxicity was found against HL-60 cells (IC50 3.1 IU/mL), which was comparable to commercial asparaginase. Apoptosis was observed in HL-60 cells treated with purified asparaginase as obtained by cell and nuclear morphological studies. Analysis of cell cycle progression indicated that apoptosis was induced by enzyme by cell cycle arrest in G0/G1 phase. Further investigation of mechanism of apoptosis by measuring mitochondrial membrane potential revealed that enzyme triggers the mitochondrial pathway of apoptosis dysfunctioning of mitochondrial integrity. The enzyme was found to be nontoxic for human noncancerous cells and nonhemolytic for human erythrocytes (Husain et al., 2016).

2.5 L-Glutaminase In most cancer, glutamine acts as primary mitochondrial substrate. Although the consumption of glutamine by cancer cells under hypoxic conditions is 15 times higher than that of normal cells, they are unable of producing their own glutamine de novo but normal cells can do so. L-Glutaminase is an amido-hydrolase enzyme that reduces blood glutamine levels by cleaving glutamine into glutamic acid and ammonia and thereby inhibits cancer cell proliferation through glutamine deprivation (Wise & Thompson, 2010). An intracellular L-glutaminase was purified from Penicillium brevicompactum NRC 829. The purified enzyme inhibited the growth of human hepatocellular carcinoma (Hep-G2), with an IC50 value of 63.3 μg/mL (Elshafei et al., 2014). L-Glutaminase produced by Alcaligenes faecalis KLU102, isolated from the marine realm (Bay of Bengal), was tested for its anticancer potency. Purified L-glutaminase exhibited dose-dependent cytotoxicity against HeLa cells, as assayed by the MTT assay, with an IC50 value of 12.5 μg/mL (Pandian et al., 2014). To enhance stability and bioavailability of the enzyme, L-glutaminase was immobilized on polyethyleneglycol

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(PEG)–polyhydroxybutyrate (PHB) nanoparticles to obtain enzymeconjugated nanoparticles of size 20–30 nm. Purified marine L-glutaminase and PEG–PHB-conjugated glutaminase nanoparticles were studied for their anticancer activity using HeLa cells. The inhibitory concentration of L-glutaminase was found to be 2.5 μg/mL. 3.5-Fold decreased DNA proliferation was witnessed in treatment group compared to control group. The enzyme was found to inhibit the proliferation of cancer cells by glutamine deprivation-induced formation of apoptotic bodies (Pandian, Deepak, Nellaiah, & Sundar, 2015).

2.6 Protease Proteases are ubiquitous enzymes that catalyze the cleavage of peptide bonds in proteins that are essential for cell growth and differentiation. A protease has been reported to have anticancer activity at a relatively low dose in comparison to an approved agent mitomycin C, being sourced from Bacillus species present in marine water. This protease was found to exhibit cytotoxicity against A549 (colon cancer) cell lines and exhibited dose-dependent cell viability with 57.6% cell viability at 50 μg toward A549 cell lines (Vijayasurya et al., 2014).

2.7 Lysozyme Lysozyme, chemically 1,4-β-N-acetylmuramidase, possesses angiogenic inhibition and antitumor activity. It functions by enhancing phagocytic action of polymorphonuclear leukocytes and macrophages, stimulating proliferation of these cells, and inhibiting the growth of malignant tumor. Marine-derived lysozyme extracted from marine invertebrates exhibits special activity compared to vertebrate lysozyme since marine invertebrates are able to sustain their activity even at low temperature, hypoxia, and high-pressure environment. The antitumor and antiangiogenesis effects of novel lysozyme obtained from marine bacillus were studied both in vitro and in vivo by Ye et al. (2008). Marine-derived lysozyme specifically inhibited the proliferation of endothelial cells (ECV304) induced by basic fibroblast growth factor (bFGF) in concentration-dependent manner (IC50 3.64 μM), while hen egg lysozyme showed no significant inhibitory effect on endothelial cells. Authors have suggested that the inhibitory effect of marine lysozyme on endothelial cells may be related with bFGF pathway as bFGF is a well-known potential endothelial cell mitogen. Moreover, this lysozyme had no cytotoxic effects on nonendothelial cell lines

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(hepatoma 22, NIH3T3, and S180), confirming its specific inhibitory action on endothelial cells. This marine lysozyme was found to suppress neovascularization in chicken embryos as assessed by chorioallantoic membrane (ChAM) assay which confirmed its antiangiogenesis activity. In vivo subcutaneous administration of marine lysozyme in mice bearing either sarcoma 180 or hepatoma 22 revealed marked tumor growth inhibition, whereas the tumor growth inhibition by hen egg lysozyme was not significant (Ye et al., 2008). Wang et al. (2009) suggested a probable mechanism of antiangiogenesis effect of marine low-temperature lysozyme (MLTL) obtained from a marine bacillus through fermentation. In an in vivo study using zebra fish embryo to assess antiangiogenic effect, treatment with MLTL substantially inhibited the growth of subintestinal vessels (SIVs) in a dose-dependent manner, and that dose of 400 μg/mL MLTL was sufficient to block the growth of SIVs. MLTL suppressed various stages of angiogenesis process, viz., the proliferation, migration, and tube formation of human umbilical vein endothelial cells (HUVECs) when studied in vitro in a dose-dependent manner. Interestingly, MLTL was found to induce apoptosis in HUVECs as confirmed by flow cytometry and DNA electrophoresis. MLTL was found to increase the intracellular concentration of Ca2+ in HUVECs which is indicative of the fact that disruption of intracellular Ca2+ homeostasis plays an influential role in induction of apoptosis of HUVECs. These findings demonstrated that MLTL inhibits angiogenesis through its pleiotropic effects on vascular endothelial cells and induction of apoptosis in endothelia cells, thereby proposing MLTL to be a promising antiangiogenic agent for cancer therapy (Wang et al., 2009).

2.8 Acetylcholinesterase Many marine-derived natural products have demonstrated anticancer activity by blocking the formation of new blood vessels (angiogenesis) and thereby inhibiting the growth of tumor. The antiangiogenic activity of acetylcholinesterase present in marine sponges was explored as a potential antitumor agent. The marine sponges Spirastrella pachyspira, Halichondria glabrata, and Cliona lobata were tested for specific acetylcholinesterase activity. S. pachyspira exhibited maximum specific enzyme activity and was further investigated for its antiangiogenic activity by ChAM assay. Antiangiogenic response of S. pachyspira extract was observed at dose of 62.5 μg/mL. At doses beyond 250 μg/mL, the S. pachyspira extract was found to be toxic,

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affecting the microvasculature of ChAM as well as normal growth and development of the embryos (Kumar & Gopalkrishnan, 2014).

2.9 Laccase Cerrena unicolor is a novel source of highly active extracellular laccase (ex-LAC) that is currently used in industry for biodegradation, bioremediation, delignification, and decolorization. Laccase belongs to group of blue multicopper oxidases that has distinctive redox ability of copper ions. The antileukemic activity of ex-LAC purified from fungi C. unicolor was first tested by Matuszewska et al. The cytotoxic effect of ex-LAC was demonstrated on human acute promyelocytic leukemia (HL-60), Jurkat, human multiple myeloma (RPMI 8226), and human chronic myeloid leukemia (K562) cell lines, as well as chronic lymphocytic leukemia (CLL) primary cells with IC50 values ranging from 0.4 to 1.1 μg/mL. The authors also reported that the obtained IC50 values of C. unicolor ex-LAC were lower in comparison with other compounds of fungal origin, thereby implying high antimalignant activity of this enzyme. Additionally, apoptotic changes in Jurkat and RPMI 8226 cells on treatment with ex-LAC when compared with control cells were evident by fluorescence microscopy and apoptosis analysis. Thus laccase was able to exert significant antileukemic activity by inducing cell apoptosis. Authors have suggested laccase to be novel therapeutic agent for the treatment of various hematological neoplasms (Matuszewska et al., 2016).

2.10 Fucoidanase Manivasagan and Oh (2015) utilized marine actinobacterium Streptomyces species as a source for producing fucoidanase. This enzyme was applied for the biosynthesis of gold nanoparticles to form fucoidanase-capped gold nanoparticles. These nanoparticles exhibited cytotoxicity against HeLa cells in a dose-dependent manner, and IC50 was found to be 350 μg/mL at 24 h and 250 μg/mL at 48 h. Analysis of nuclear morphology further revealed characteristic apoptotic changes in the HeLa cells such as chromatin condensation, fragmentation of the nucleus, and formation of apoptotic bodies. Based on these observations, authors have suggested induction of apoptosis as a possible mechanism for the antiproliferative activity of fucoidanasecapped nanoparticles (Manivasagan & Oh, 2015).

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R.H. Prabhu et al.

3. CONCLUSION Marine life holds great promise as a largely untapped source of biologically active compounds with pharmacological potential. Marine organisms are considered as important biofactory of novel enzymes due to their chemical, biological, and ecological diversity. Marine organisms have attracted special attention recently for their ability to produce bioactive enzymes. Many marine enzymes have been identified with promising anticancer activity in vitro. Further animal model studies are required to authenticate use of such marine-derived enzymes as cancer therapeutic agent. A great deal of research is required in order to realize the application of marine-sourced enzymes in clinics. Enzymes in combination with drugs would induce synergistic anticancer effect and enable reduction in side effects associated with drugs.

ACKNOWLEDGMENTS R.H.P. is thankful to Department of Science and Technology, Government of India for providing senior research fellowship under the INSPIRE scheme.

REFERENCES Bakunina, I. Y., Nedashkovskaya, O. I., Alekseeva, S. A., Ivanova, E. P., Romanenko, L. A., Gorshkova, N. M., … Mikhailov, V. V. (2002). Degradation of fucoidan by the marine proteobacterium Pseudoalteromonas citrea. Microbiology, 71, 41–47. Basha, N. S., Rekha, R., Komala, M., & Ruby, S. (2009). Production of extracellular antileukaemic enzyme L-asparaginase from marine actinomycetes by solid-state and submerged fermentation: Purification and characterisation. Tropical Journal of Pharmaceutical Research, 8, 353–360. Bowles, T. L., Kim, R., Galante, J., Parsons, C. M., Virudachalam, S., Kung, H. J., & Bold, R. J. (2008). Pancreatic cancer cell lines deficient in argininosuccinate synthetase are sensitive to arginine deprivation by arginine deiminase. International Journal of Cancer, 123, 1950–1955. Crawford, A. D., Jaspars, M., De Pascale, D., Andersen, J. H., Reyes, F., & Ianora, A. (2016). The marine biodiscovery pipeline and ocean medicines of tomorrow. Journal of the Marine Biological Association of the United Kingdom, 96, 151–158. Demain, A. L., & Sanchez, S. (2009). Microbial drug discovery: 80 years of progress. The Journal of Antibiotics, 62, 5–16. Dhevagi, P., & Poorani, E. (2006). Isolation and characterization of L-asparaginase from marine actinomycetes. Indian Journal of Biotechnology, 5, 514–520. El-Sayed, A. S., Hassan, M. N., & Nada, H. (2015). Purification, immobilization, and biochemical characterization of L-arginine deiminase from thermophilic Aspergillus fumigatus KJ434941: Anticancer activity in vitro. Biotechnology Progress, 31, 396–405. El-Sayed, A. S., Shindia, A. A., Diab, A. A., & Rady, A. M. (2014). Purification and immobilization of L-arginase from thermotolerant Penicillium chrysogenum KJ185377.1; with

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unique kinetic properties as thermostable anticancer enzyme. Archives of Pharmacal Research, 1–10. Elshafei, A. M., Hassan, M. M., Ali, N. H., Abouzeid, M. A. E., Mahmoud, D. A., & Elghonemy, D. H. (2014). Purification, kinetic properties and antitumor activity of l-glutaminase from Penicillium brevicompactum NRC 829. British Microbiology Research Journal, 4(1), 97–115. Gupta, A. P., Pandotra, P., Sharma, R., Kushwaha, M., & Gupta, S. (2013). Marine resource: A promising future for anticancer drugs. In A. U. Rahman (Ed.), Studies in natural products chemistry (pp. 229–325). Amsterdam: Elsevier. Husain, I., Sharma, A., Kumar, S., & Malik, F. (2016). Purification and characterization of glutaminase free asparaginase from Enterobacter cloacae: In-vitro evaluation of cytotoxic potential against human myeloid leukemia HL-60 cells. PLoS One, 11, e0148877. http://dx.doi.org/10.1371/journal.pone.0148877. Jeon, Y. J., Samarakoon, K. W., & Elvitigala, D. A. (2015). Marine-derived pharmaceuticals and future prospects. In S. K. Kim (Ed.), Springer handbook of marine biotechnology (pp. 957–968). Berlin, Heidelberg: Springer. Kerr, R. G., & Kerr, S. S. (1999). Marine natural products as therapeutic agents. Expert Opinion on Therapeutic Patents, 9, 1207–1222. Kiruthika, J., & Saraswathy, N. (2013). Production of L-glutaminase and its optimization from a novel marine isolate Vibrio azureus JK-79. African Journal of Biotechnology, 12, 6944–6953. Kumar, M. S., & Gopalkrishnan, S. (2014). Evaluation, partial characterization and purification of acetylcholine esterase enzyme and antiangiogenic activity from marine sponges. Journal of Coastal Life Medicine, 2, 849–854. Kumari, P., Sankar, G., & Prabhakar, T. (2011). L-Asparaginase production and molecular identification of marine streptomycete strain WS3/1. International Journal of Pharmacy and Biomedical Research, 2, 244–249. Manivasagan, P., & Oh, J. (2015). Production of a novel fucoidanase for the green synthesis of gold nanoparticles by Streptomyces sp. and its cytotoxic effect on HeLa cells. Marine Drugs, 13, 6818–6837. Manna, S., Sinha, A., Sadhukhan, R., & Chakrabarty, S. L. (1995). Purification, characterization and antitumor activity of L-asparaginase isolated from Pseudomonas stutzeri MB-405. Current Microbiology, 30, 291–298. Martins, A., Vieira, H., Gaspar, H., & Santos, S. (2014). Marketed marine natural products in the pharmaceutical and cosmeceutical industries: Tips for success. Marine Drugs, 12, 1066–1101. Matuszewska, A., Karp, M., Jaszek, M., Janusz, G., Osi nska-Jaroszuk, M., Sulej, J., & Giannopoulos, K. (2016). Laccase purified from Cerrena unicolor exerts antitumor activity against leukemic cells. Oncology Letters, 11, 2009–2018. Mayer, A. M., Glaser, K. B., Cuevas, C., Jacobs, R. S., Kem, W., Little, R. D., … Shuster, D. E. (2010). The odyssey of marine pharmaceuticals: A current pipeline perspective. Trends in Pharmacological Sciences, 31, 255–265. Newman, D. J., & Cragg, G. M. (2016). Drugs and drug candidates from marine sources: An assessment of the current “state of play” Planta Medica, 82, 775–789. Pandian, S. R. K., Deepak, V., Nellaiah, H., & Sundar, K. (2015). PEG–PHB-glutaminase nanoparticle inhibits cancer cell proliferation in vitro through glutamine deprivation. In Vitro Cellular & Developmental Biology Animal, 51, 372–380. Pandian, S. R. K., Deepak, V., Sivasubramaniam, S. D., Nellaiah, H., & Sundar, K. (2014). Optimization and purification of anticancer enzyme L-glutaminase from Alcaligenes faecalis KLU102. Biologia, 69, 1644–1651. Philip, R., Campbell, E., & Wheatley, D. N. (2003). Arginine deprivation, growth inhibition and tumour cell death: 2. Enzymatic degradation of arginine in normal and malignant cell cultures. British Journal of Cancer, 88, 613–623.

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Prabhu, R. H., Patravale, V. B., & Joshi, M. D. (2015). Polymeric nanoparticles for targeted treatment in oncology: Current insights. International Journal of Nanomedicine, 10, 1001–1018. Savitri, A. N., & Azmi, W. (2003). Microbial L-asparaginase: A potent antitumour enzyme. Indian Journal of Biotechnology, 2, 184–194. Saxena, S. (2015). Microbial enzymes and their industrial applications. In Saxena S. (Ed.), Applied microbiology (pp. 121–154). New Delhi: Springer India. Schwartsmann, G., da Rocha, A. B., Mattei, J., & Lopes, R. (2003). Marine-derived anticancer agents in clinical trials. Expert Opinion on Investigational Drugs, 12, 1367–1383. Selvam, K., & Vishnupriya, B. (2013). Partial purification and cytotoxic activity of L-asparaginase from Streptomyces acrimycini NGP. International Journal of Research in Pharmaceutical and Biomedical Sciences, 4, 859–869. Senthil, K. M., Selvam, K., & Singaravel, R. (2012). Homology modeling of a unique extracellular glutaminase free l-asparaginase from novel marine actinomycetes. International Journal of Research in Biotechnology and Biochemistry, 2, 1–12. Shakambari, G., Birendranarayan, A. K., Lincy, M. J. A., Rai, S. K., Ahamed, Q. T., Ashokkumar, B., … Varalakshmi, P. (2016). Hemocompatible glutaminase free L-asparaginase from marine Bacillus tequilensis PV9W with anticancer potential modulating p53 expression. RSC Advances, 6, 25943–25951. Suleria, H. A. R., Osborne, S., Masci, P., & Gobe, G. (2015). Marine-based nutraceuticals: An innovative trend in the food and supplement industries. Marine Drugs, 13, 6336–6351. Unissa, R., Sudhakar, M., & Reddy, A. S. K. (2015). Selective isolation and molecular identification of L-arginase producing bacteria from marine sediments. World Journal of Pharmacy and Pharmaceutical Sciences, 4, 998–1006. Unissa, R., Sudhakar, M., & Reddy, A. S. K. (2016). Evaluation of in vitro anti-proliferative activity of L-arginine deiminase from novel marine bacterial isolate. British Microbiology Research Journal, 13, 1–10. Vijayasurya, G. D. M., Tintu, S. P., & Manjusha, W. A. (2014). In vitro cytotoxicity and antimicrobial screening of protease enzyme isolated from marine bacteria. Indian Journal of Scientific Research and Technology, 2, 18–22. Wang, Z., Liu, J., Su, A., Sun, M., & Wang, C. (2009). Antiangiogenic activity of lowtemperature lysozyme from a marine bacterium in vivo and in vitro. Chinese Journal of Oceanology and Limnology, 27, 835–844. Wise, D. R., & Thompson, C. B. (2010). Glutamine addiction: A new therapeutic target in cancer. Trends in Biochemical Sciences, 35, 427–433. Ye, J., Wang, C., Chen, X., Guo, S., & Sun, M. (2008). Marine lysozyme from a marine bacterium that inhibits angiogenesis and tumor growth. Applied Microbiology and Biotechnology, 77, 1261–1267.

CHAPTER TWO

Bacillus Probiotic Enzymes: External Auxiliary Apparatus to Avoid Digestive Deficiencies, Water Pollution, Diseases, and Economic Problems in Marine Cultivated Animals Jorge Olmos Soto1 Molecular Microbiology Laboratory, Centro de Investigacio´n Cientı´fica y de Educacio´n Superior de Ensenada (CICESE), Ensenada, Baja California, Mexico 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Commercial Diet Ingredients Preferentially Utilized 2.1 Fish Meal and Fish Oil as Main Ingredients 2.2 Land-Vegetable Resources 3. Enzyme Deficiencies in Cultivated Marine Animals 4. Marine Animal Diseases Induced by ANFs 5. Water Pollution and Economic Problems as a Consequence of Poor Digestion– Assimilation Process 6. Bacillus as the Enzyme Production Machinery 7. B. subtilis Probiotic Bacterium 8. Conclusions Acknowledgment References

16 18 20 21 26 27 27 28 29 31 32 32

Abstract Exploitation of marine fishes is the main source of several life-supporting feed compounds such as proteins, lipids, and carbohydrates that maintain the production of most trading marine organisms by aquaculture. However, at this rate the marine inventory will go to the end soon, since fishery resources are finite. In this sense, the availability of the principal ingredients obtained from marine fishes is going to decrease considerably, increasing the diet prices and affecting the economy of this activity. Therefore, aquaculture industry needs to find nonexpensive land unconventional resources of protein, carbohydrates, and lipids and use bacterial probiotics to improve Advances in Food and Nutrition Research, Volume 80 ISSN 1043-4526 http://dx.doi.org/10.1016/bs.afnr.2016.11.001

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2017 Elsevier Inc. All rights reserved.

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Jorge Olmos Soto

digestion–assimilation of these unfamiliar compounds. Bacillus subtilis is a cosmopolitan probiotic bacterium with a great enzymatic profile that could improve nutrient digestion–assimilation, induce healthy growth, and avoid water pollution, decreasing economic problems and increasing yields in the aquaculture industry. In this chapter, we present how Bacillus enzymes can help marine animals to assimilate nutrients from unconventional and economic plant resources.

1. INTRODUCTION Seafood is a tradition in many countries and in terms of health benefits has an excellent nutritional profile. It is a good source of protein, lipids, vitamins, minerals, and essential micronutrients; however, fisheries capture has been declining for the last 20 years (Klinger & Naylor, 2012; Tacon & Metian, 2008). Aquaculture is practiced by both poor farmers and multinational companies around the world. Industrial production of fishes and crustaceans has a significant social, environmental, and economic impact on countries involved in this activity. In this sense, aquaculture is one of the fastest-expanding agricultural industries in the world, with growth rates exceeding 10% each year (FAO, 2016; Yajie & Sumaila, 2008). Nevertheless, main ingredients used to cultivate marine animals are in a dangerous limit because of the overexploitation of the marine fishery resources. The state of wild marine resources raises concern as, since 1990, about a quarter of them are seriously overfished producing great environmental impact all over the world, inducing ecological and economic problems (FAO, 2016). In this sense, some authors have predicted that inclusion of fish meal and fish oil in aquaculture diets will be minimum or null in 2020 (Tacon & Metian, 2008). Among the most harvested species are anchovy, herring, sardines, mackerel, and krill, and fish meal and fish oil for aquaculture diets are obtained principally from these species. As the aquaculture industry increases its demand for fish meal and fish oil, the overexploitation of fishery resources increases too. In fact, 7 of the world’s top 10 fisheries are dedicated to produce fish meal and fish oil for the aquaculture industry (http://www. worldwildlife.org/industries/fishmeal-and-fish-oil). From a ton of processed fish approximately 25% of fishmeal is obtained, containing an average of 60% digestible protein. Regarding fish oil the amount generated by a ton of processed fish is around 10%. Moreover, fish oil and fish meal requirements for cultivated marine species like shrimp and salmon are between 10% and 50%, which increases the cost of aquaculture diets (Table 1, Fig. 1).

Bacillus Probiotic Enzymes

17

Table 1 Production of Economically Important Cultivated Species (FAO, 2016) Common Name Binomial Name Harvest (tons)

Grass carp

Ctenopharyngodon idellus

5,537,794

Silver carp

Hypophthalmichthys molitrix

4,967,739

Common carp

Cyprinus carpio

4,159,117

Nile tilapia

Oreochromis niloticus

3,670,260

Whiteleg shrimp

Penaeus vannamei

3,668,682

Tiger shrimp

Penaeus monodon

634,522

Atlantic salmon

Salmo salar

2,326,288

Milkfish

Chanos chanos

1,039,184

Rainbow trout

Oncorhynchus mykiss

812,940

Amur catfish

Silurus asotus

455,791

Swamp eel

Monopterus albus

358,036

In fish and crustaceous aquaculture, feeds represent the most expensive production cost (50–70%). In this sense, the quality of diets is an important factor to increase nutrient assimilation and growth, to improve animal health, and to avoid disease proliferation. For these reasons, there is an increasing interest in developing digestible, safe, economic, and environmentally friendly formulations (Olmos, Paniagua-Michel, Lo´pez, & Ochoa, 2014). Salmon is one of the prized marine animals; however, the wild Atlantic salmon fishery is commercially dead after extensive habitat damage and overfishing. Wild fisheries produce only 0.5% of the Atlantic salmon available in world fish markets, and the rest is predominantly aquacultured in Norway, Chile, United Kingdom, and Canada (Liu, Chuenpagdee, & Sumalia, 2013). Research based on the experimental diets showed that juvenile Atlantic salmon reared in seawater required 50% of protein and approximately 20% of lipids and tolerate no more than 10% of carbohydrates (Einen & Roem, 1997). However, optimum protein and lipid-level inclusion in marine animals’ feeds depend upon the dietary energy available derived from an adequate carbohydrates’ digestion–assimilation process (Arellano-Carbajal & Olmos-Soto, 2002; Lopez et al., 2016; Olmos, Ochoa, Paniagua-Michel, & Contreras, 2011). On the other hand, commercial diets produced for shrimp are made approximately with 20–30% of protein, 50–60% of carbohydrates, and

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5–10% of total lipids (Olmos et al., 2011). Carbohydrates can substitute proteins as energy source in fish and shrimp feeds due to their natural constitution (Ochoa, Paniagua-Michel, & Olmos, 2014). However, this marine carnivorous also have severe restrictions for digestion–assimilation of carbohydrates from the vegetable origin (Arellano-Carbajal & Olmos-Soto, 2002; Le Chevalier & Van Wormhoudt, 1998). Therefore, functional feeds (FF) development will represent one of the greatest advances and improvements in the aquaculture industry, promoting healthy growth of the cultivated animals, improving their immune system, and inducing beneficial physiological effects (Olmos et al., 2014). Thus, using land-vegetable ingredients and probiotic bacteria like Bacillus subtilis, FF will bring the opportunity to reduce or eliminate fish meal and fish oil from diets and to make the activity more profitable (Ochoa-Solano & Olmos, 2006).

2. COMMERCIAL DIET INGREDIENTS PREFERENTIALLY UTILIZED There are around 40 essential nutrients needed by all animals including vitamins, dietary minerals, essential fatty acids, and essential amino acids. In this chapter, we will focus more specifically on proteins, carbohydrates, and lipids, and on their optimal digestion–assimilation processes. Marine most trading species need diets containing high levels of proteins (amino acids), carbohydrates (energy), lipids (fatty acids and omegas), and micronutrients (vitamins, minerals, and pigments). In aquaculture, artificial diets supply all the needs for an optimal healthy growth of the marine species; however, traditional feeds are made from fish meal and fish oil as the main ingredients (Olmos et al., 2014). Several decades ago, small pelagic fishes were the greatest economical option for protein and oil sources, but the overexploitation of fishery resources caused ecological imbalance and increased diet prices. Thus, it is predicted that by 2020 the amount of fish meal and fish oil in diets for aquaculture will be minimal or null (Klinger & Naylor, 2012; Tacon & Metian, 2008). Today, the commercial feeds used for carnivorous fishes and crustaceans contain 50% and 30% of protein, 10% and 40% of carbohydrates, and 20% and 4% of lipids, respectively (Lopez et al., 2016; Olmos et al., 2011; Table 2). Carnivorous marine fishes “tolerate” carbohydrate concentrations no greater than 10% and “need” 50% of protein for optimal growth. Due to the low capacity of marine carnivorous fishes to digest–assimilate carbohydrates, these animals obtain their energy requirements directly from

Table 2 Nutrient Requirements for Atlantic Salmon (Lall, 2008) Stage/Size Class Nutrients

Fry

Fingerling

Juvenile

Grower

Marine Grower

Broodstock

Crude protein, % min

50

45–50

45

42–45

40–45

45

Arginine

2

2

2

1.6

1.6

1.6

Histidine

0.7

0.7

0.7

0.8

0.7

0.7

Isoleucine

0.8

0.8

0.8

0.8

0.8

0.8

Leucine

1.4

1.4

1.4

1.4

1.4

1.4

Lysyne

2

1.8

1.8

1.8

1.8

1.8

Methionine

1.1

1

1

1

1

1

Phenylalanine

1.2

1.2

1.2

1.2

1.2

1.2

Threonine

0.8

0.8

0.8

0.8

0.8

0.8

Tryptophan

0.2

0.2

0.2

0.2

0.2

0.2

Valine

1.3

1.3

1.3

1.3

1.3

1.3

Carbohydrate, % max

10

10

12

12

12

12

Crude fiber, % max

2

3

3

3

3

3

Digestible energy, min kJ/g

19

19

19

20

20

19

Protein to energy ratio, mg/kJ

23–24

22–23

21–22

20–21

17–18

18

Crude lipid, % min

16–18

20

20

20–24

24–30

24

Amino acids, % min of dietary protein

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Jorge Olmos Soto

proteins, increasing the diet prices (Buddington, Buddington, Deng, Hemre, & Wilson, 2002; Lopez et al., 2016). Even when shrimps have the highest threshold than carnivorous fishes to tolerate complex carbohydrates, their incapacity to digest several types of these compounds still remains (Ochoa et al., 2014; Olmos et al., 2011). Regarding this information, the aquaculture industry needs to find land unconventional nutrient sources that satisfy the marine animal requirements. An alternative is to use vegetable ingredients because they contain proteins, carbohydrates, and lipids capable to substitute those present in fish meal and fish oil. However, assimilation of vegetable ingredients is a big concern because the presence of antinutritional factors (ANFs) (Olmos et al., 2014; Refstie, Sahlstrfmc, Brathenc, Baeverfjorda, & Krogedald, 2005). High levels of ANFs cannot be tolerated by aquacultured animals because of their enzyme deficiencies or by the presence of enzyme inhibitor (Sørensen et al., 2011).

2.1 Fish Meal and Fish Oil as Main Ingredients Fish meal and fish oil are natural ingredients of high nutritional value and are the main ingredients in aquaculture feeds because they supply essential amino acids and lipids needed for an animal’s development. Fish meal is a source of high-quality protein with a well-balanced essential and nonessential amino acids’ profile (Tacon & Metian, 2008). In view of its advantageous characteristics, aquaculture has continued to increase the demand for fish products during the last 50 years. Additionally, fish meal and fish oil provide a balanced amount of the most essential minerals, phospholipids, and fatty acids, which help to increase the growth rate and production yields. Fish oil is a great source of omega-3 eicosapentaenoic fatty acid (EPA) and docosahexaenoic fatty acid, which are not made by the fish but become concentrated further up the food chain from the marine phytoplankton. However, fish meal and fish oil are generally expensive and are not always available. In addition, it is well documented that the marine resources from fisheries capture are going to the end soon. In this sense, fish meal and fish oil prices have been increasing exponentially during the last two decades, reducing profits from aquaculture (Klinger & Naylor, 2012; Tacon & Metian, 2008; Fig. 1). Many ingredients of animal and vegetable origin have been used to substitute fish meal and fish oil without the expected success (Enes, Panserat, Kaushik, & Oliva-Teles, 2008; German, Horn, & Gawlicka, 2004; Hemre, Shiau, Deng, Storebakken, & Hung, 2002; Samocha, Davis, Saoud, & DeBault, 2004; Sookying & Davis, 2011). Another attempt was

Bacillus Probiotic Enzymes

21

Fig. 1 Fish meal and fish oil prices.

made trying to adapt diet ingredients to the enzymatic profile of the animals; however, this strategy has not been practical or economically viable for the aquaculture industry. Using proteins, carbohydrates, and lipids from the vegetable origin could improve yields of the activity because the technology to produce grains from agriculture is getting better each year.

2.2 Land-Vegetable Resources Grains like soy, corn, wheat, and sorghum constitute a great source of proteins, carbohydrates, and lipids. Their production has been increasing each year, maintaining competitive prices; therefore the utilization of these compounds could be a solution for the aquaculture industry. However, marine fishes and crustaceous are incapable to digest high levels of carbohydrates and proteins from the land origin, due to enzyme deficiencies in the animals and the presence of enzyme inhibitors in the grains (Cruz-Suarez, Ricque, Pinal-Mansilla, & Wesche-Ebelling, 1994; Dall, Hill, Rothlisberg, & Sharples, 1990; Olmos et al., 2011; Sørensen et al., 2011). Nowadays, one important goal is to reduce fish meal and fish oil used in aquaculture feeds, while human health benefits of seafood consumption are maintained. Carnivorous fishes such as salmon tolerate no more than 10% of carbohydrates, which prevents the inclusion of these compounds in marine fish diets at high concentrations (Lopez et al., 2016). However, the inability of carnivorous fishes to tolerate high levels of carbohydrates does not mean that these molecules could not be assimilated and utilized by fish cells, once they were previously digested in sugars. Additionally, vegetable proteins could be the solution to improve profits and the economy of aquaculture

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Jorge Olmos Soto

industry due to its low prices in comparison with fish protein. However, enzyme inhibitors contained in grains reduce considerably the digestion– assimilation of these molecules (Sørensen et al., 2011). In this sense, the application of probiotic bacteria with a great level and diversity of proteases, carbohydrases, and lipases production could be a nonexpensive and healthy solution to improve the profits of aquaculture (Olmos et al., 2014). 2.2.1 Soy Protein Soybean contains 40% of protein of the highest quality, which is almost equal to that of meat and milk, because it contains essential amino acids in the amounts needed by the animals and humans (Jenkins et al., 2010). This makes soybean protein the greatest nutritional source compared with other beans and legumes (Table 3; Fig. 2). 2.2.2 Soy Carbohydrates Supposedly, carbohydrates are the easiest assimilable source of energy; most of them could be converted into glucose during digestion and be assimilated Table 3 Grain Nutrients’ Content Grains Macromolecule

Approx. %

Soy

Protein

40

Carbohydrate

30

Lipids

20

Other compounds and water

10

Protein

10

Carbohydrate

70

Lipids

10

Other compounds and water

10

Protein

16

Carbohydrate

72

Lipids

2.0

Other compounds and water

10

Protein

12

Carbohydrate

72

Lipids

4.0

Other compounds and water

12

Corn

Wheat

Sorghum

Bacillus Probiotic Enzymes

23

Fig. 2 Crystal structure of legumin.

in the small intestine (Ochoa et al., 2014; Stone, 2003). Soybean contains 30% of carbohydrates, of which 7% are sugars, 9% is fiber, and 14% belongs to sucrose (8%), raffinose (2%), and stachyose (4%). In this sense, some of the soybean carbohydrates are nondigestible and toxic for the monogastric animals like carnivorous fishes cultivated in aquaculture (Choct, Dersjant-Li, McLeish, & Peisker, 2010). 2.2.3 Soy Lipids Approximately 20% of the calories in soybeans are derived from lipids (Fig. 3): polyunsaturated 12% (omega-6 and omega-3), monounsaturated 5% (omega-9), and saturated 3% (primarily palmitic acid). The polyunsaturated fatty acids contained in soybeans are of great interest because they are similar to those present in the fish oil used to prepare commercial feeds in aquaculture. 2.2.4 Soy Antinutritional Factors There are a great number of ANFs present in soybean; among those factors are the protease inhibitors that induce pancreatic hypertrophy/hyperplasia, which results in an inhibition of growth and death (Dipietro & Liener, 1989). Additionally, lectins could bind to glycoprotein receptors on the epithelial cells lining the intestinal mucosa and inhibiting growth by interfering with the absorption of nutrients. Other ANFs are the previously mentioned

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Jorge Olmos Soto

Soy lipid Saturated fat 15%

Omega-3 8%

Omega-9 23%

Omega-6 54%

Fig. 3 Lipid content in soybeans.

nondigestible oligosaccharides that become toxic molecules due to enzyme deficiencies in animals. Soy ANFs could induce allergenic responses in humans as well as in calves, piglets, and fishes. Some of these compounds inhibit the utilization of soy protein by monogastric animals, like the marine carnivorous fishes and crustaceous cultivated in aquaculture (Liener, 1994; Sørensen et al., 2011). In this sense, attempts to purify soy protein had been done, but prices make the product economically unviable to be utilized in this activity. 2.2.5 Other Grains 2.2.5.1 Carbohydrates

Aside from containing a variable amount of water and fiber, corn, wheat, and sorghum are mainly composed by carbohydrates (Table 3). The most abundant carbohydrate in these grains is starch that contains 25% of amylose and 75% of amylopectin, which are constituted by glucose molecules in either occasionally branched chains or unbranched chains (Ochoa et al., 2014; Fig. 4). Corn, wheat, and sorghum contain around 70% of starch that could be a great source of energy in aquaculture feeds; however, marine carnivorous fishes and shrimps do not produce α-1,6-glucosidases to digest α-1,6-bonds from amylopectin branches (Krogdahl, Hemre, & Mommsen, 2005; Le Chevalier & Van Wormhoudt, 1998). In this sense, the amylopectin

Bacillus Probiotic Enzymes

25

A

B CH2OH

CH2OH

O

O

OH CH2OH

CH2OH

O

CH2OH

O

OH OH

O OH

OH

OH

O OH

OH

O

OH

OH

CH2OH

O OH

OH x

OH

O CH2

O

CH2OH O

OH

O

OH

OH

OH

O OH

O OH

OH x

OH

Fig. 4 Starch polymers constituted by glucose molecules: (A) amylose and (B) amylopectin.

molecule could be considered as ANF to these animals—the reason why fishes tolerate no more than 10% of starch in their diet. In addition, shrimps also lack α-1,6-glucosidase enzyme; nevertheless, commercial diets contain until 50% of starch inclusion (Arellano-Carbajal & Olmos-Soto, 2002; Olmos et al., 2011). Nondigested amylopectin could produce several health problems induced by intoxication and allergic reactions. Moreover, nondigested starch could be an important source of water pollution and disease proliferation. In this sense, the utilization of probiotic bacteria with the capacity to produce α-1,6-glucosidases could improve starch degradation and assimilation. Additionally, available energy generated by the digestion of starch could diminish the diet prices because protein consumption will be derived exclusively to the animal growth and will not be wasted as energy source (Olmos et al., 2014). 2.2.5.2 Proteins and Lipids

The percentage of protein contained in corn, wheat, and sorghum is similar (Table 3); nevertheless, gluten present in wheat is highly toxic for humans with celiac disease (Moro´n et al., 2008). In addition, sorghum protein is considered of low quality due its amino acid constitution and some inhibitors (Chandrasekher, Suryaprasad, & Thillaisthanam, 1981). For these reasons, none of these proteins have been included at high levels in aquaculture diets. In this sense, probiotics could help to digest and utilize these kinds of proteins; however, everything depends on the selected bacteria. With respect to lipids, these grains produce high levels of omega-6 polyunsaturated fatty acid but low levels of omega-3, a disadvantageous ratio with respect to fish oil and soy lipids. In this sense, other economical alternative for omega-3

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Jorge Olmos Soto

H

J J

H

J J

H

J J

H

J J

H

J J

H

H

J J

H

H

J J

H

H

J J

J J

H

J J

H

J J

H

HJ CJ C JCJ C J C J C J C J C J CJ CJ C J C H

H

H

H

H

H

H

J

H

O K OH

Saturated fatty acid J

J

J

J J

J

J J

J

J J

J J

J

J

H

J J

H O K C C C J J H H C JC JC K H H OH H C JC J C K HJ C J CJ C K H H H H H H H H

H

Unsaturated fatty acid

Fig. 5 Saturated and unsaturated fatty acid molecules.

Corn lipid Saturated fat 13%

Omega-9 29%

Omega-3 1%

Omega-6 57%

Fig. 6 Corn lipid content.

production is found in microalgae, which in the last years have been investigated intensively for biofuel production (Hannon, Gimpel, Tran, Rasala, & Mayfield, 2010; Rubio & Olmos, 2015) (Figs. 5 and 6).

3. ENZYME DEFICIENCIES IN CULTIVATED MARINE ANIMALS Deficiency of some proteases, carbohydrases, and lipases in the marine cultivated animals is the main problem for the optimal digestion and assimilation of several vegetable compounds, limiting their high-level inclusion in

Bacillus Probiotic Enzymes

27

diets. In this sense, the restricted capacity of fishes and shrimps to digest starch causes fish meal to be utilized by animals as a source of protein and energy, making feeding less efficient and more expensive. Additionally, the incapacity of marine animals to digest some of the soybean carbohydrates transformed these molecules in ANFs, inducing severe damages and pathologies (Krogdahl, Bakke-Mckellep, & Baeverfjord, 2003). On the other hand, as the soybean protein contains ANFs like the enzyme inhibitor that reduces the possibility to supplement high levels of this ingredient in fish and shrimp diets. Since many of the marine cultivated animals lack specific intestinal proteases, carbohydrases, and lipases for the degradation of ANFs, the supplementation of these enzymes using probiotics like B. subtilis will result in a better digestion–assimilation, bigger and healthier animals, and less pollution and diseases.

4. MARINE ANIMAL DISEASES INDUCED BY ANFs Nutritional status is considered one of the most important conditions in animals to avoid and resist diseases. Outbreaks of fish and shrimp diseases commonly occur when these animals are stressed due to a variety of factors including poor nutrition. The need for an appropriate feeding to improve the health status and to prevent diseases of aquacultured animals is widely recognized (Toranzo, Magarin˜os, & Romalde, 2005). In this sense, the losses in aquaculture rise up to 60% because of the environmental pollution and diseases, both induced principally as a consequence of poorly formulated diets and the animal’s enzyme deficiencies. Therefore, the mechanisms through feeding impacts the immune system, and the relationship between diet ingredients and the susceptibility to infectious diseases must be studied. Thus, it will be possible to manipulate diets to improve the health status of the animals (Bricknell & Dalmo, 2005).

5. WATER POLLUTION AND ECONOMIC PROBLEMS AS A CONSEQUENCE OF POOR DIGESTION–ASSIMILATION PROCESS Fish and shrimp ponds’ water pollution is principally generated by excessive inputs of nonassimilated nutrients and feces from the animals (Mente, Graham, Begon˜a, & Neofitou, 2006). Ponds water eutrophication by nitrogen or phosphorus stimulates the algal growth and the

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development of pathogenic microorganisms (US Environmental Protection Agency, EPA). Algae blooms and spread of diseases in ponds can drastically increase animal losses and the remediation treatment cost. In this sense, an accurate digestion–assimilation process from most of the diet ingredients will drastically reduce problems induced by inappropriate feeding. Furthermore, profits will be increased because of the increase in products of better quality.

6. BACILLUS AS THE ENZYME PRODUCTION MACHINERY Digestion is a relevant process in animal nutrition since it influences the bioavailability and enhances the assimilation of nutrients needed for fish and shrimp growth. A poor digestion induced by complex diet ingredients known as ANFs and the deficiency of specific degradative enzymes in the animals are a bad combination to the aquaculture industry. However, vegetable ingredients added to diets are an increasing tendency; therefore, Bacillus probiotic supplementation in feeds must be carried out as an external enzymatic machinery to increase digestion–assimilation of ANFs and to improve health status (Table 4). Species of the Bacillus genus are among the most widespread microorganisms in nature; they can be found in soil, water, air, plants, animals, and humans. Bacillus constitutes a diverse group of rod-shaped Gram-positive bacteria, characterized by their ability to produce a great variety of enzymes and robust spores (Andersson, Weiss, Rainey, & Salkinoja-Salonen, 1999; Garbeva, Van Veen, & Van Elsas, 2003; Ivanova et al., 1999; Nicholson, 2004; Sonnenschein, Losick, & Hoch, 1993). In this sense and based on our own experiences, we defined probiotics from the Bacillus genus as living microbial supplement that: (a) positively affects host health status by modifying the host-associated microbial community and its immune system; (b) secretes a great variety of enzymes to increase feed degradation–assimilation enhancing its nutritional values; (c) improves the quality of environmental parameters by the bioremediation of waste products; and (d) support the extreme environmental changes (Fig. 7). Thus, the use of Bacillus species as probiotics in animal feeds is an attractive opportunity for the aquaculture industry because these bacteria meet all the requirements and some of them are recognized as safe by the FDA.

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Table 4 Bacillus Enzymes (Bron, Meima, van Dijl, Wipat, & Harwood, 1999; OchoaSolano & Olmos, 2006; Pangsri, Piwpankaew, Ingkakul, Nitisinprasert, & Keawsompong, 2015; Selim & Reda, 2015; Sumathi et al., 2014) Enzyme Producer Strain

α-Amylase

B. amyloliquefaciens, B. circulans, B. licheniformis, B. stearothermophilus, B. subtilis

β-Amylase

B. polymyxa, B. cereus, B. megaterium

Alkaline phosphatase

B. licheniformis, B. subtilis

Cyclodextran glucanotransferase

B. macerans, B. megaterium, Bacillus sp.

β-Galactosidase

B. stearothermophilus, B. subtilis

β-Glucanase

B. subtilis, B. circulans

β-Glucosidase

B. subtilis

Glucose isomerase

B. coagulans

Glucosyl transferase

B. megaterium

Glutaminase

B. subtilis

Galactomannase

B. subtilis

β-Lactamase

B. licheniformis

Lipase

Bacillus subtilis

Metalloprotease

B. lentus, B. polymyxa, B. subtilis, B. thermoproteolyticus

Neutral protease

B. amyloliquefaciens, B. subtilis

Penicillin acylase

Bacillus sp.

Pullulanase

Bacillus sp., B. acidopullulans

Serine protease

B. amyloliquefaciens, B. amylosaccharicus, B. licheniformis, B. subtilis

Urease

Bacillus sp.

Uricase

Bacillus sp.

7. B. SUBTILIS PROBIOTIC BACTERIUM The B. subtilis genome is totally sequenced, leading to the generation of a great amount of basic and applied knowledge in this bacterium. B. subtilis is not harmful to mammals including humans; hence, it is

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Stimulates immune system

Improves health status Eliminates pathogenic bacteria Bacillus subtilis

Increases feed degradation

Survives extreme conditions Induces bioremediation

Fig. 7 Bacillus subtilis probiotic capacities.

recognized as GRAS by the FDA. Additionally, this bacterium is commercially important as the producer of a great variety of secondary metabolites like antibiotics, fine chemicals, and enzymes, as well as heterologous proteins, antigens, and vaccines (Berdy, 2005; Ferrari, Jarnagin, & Schmidt, 1993; Olmos & Contreras-Flores, 2003; Sonnenschein et al., 1993; Stein, 2005; Valdez, Yepiz-Plascencia, Ricca, & Olmos, 2014; Westers, Westers, & Quax, 2004). B. subtilis grows efficiently in almost all sources of carbon and nitrogen because its enzymes are very efficient, digesting a great variety of proteins, carbohydrates, and lipids from any origin including plants (Arellano-Carbajal & Olmos-Soto, 2002; Ochoa-Solano, 2012; Ochoa-Solano & Olmos, 2006; Sonnenschein et al., 1993). The enzymes also break down organic debris from shrimp/fish cultures inducing ponds bioremediation and preventing the spread of viral and bacterial diseases (Guo et al., 2009; Olmos et al., 2011; Paniagua-Michel, Franco-Rivera, Cantera, & Stein, 2005; Rengpipat, Phianphak, Piyatiratitivorakul, & Menasvetac, 1998). Moreover, the antimicrobial activity of Bacillus species is also carried out by their ability to produce antibiotics, principally from the peptide origin (Khochamit, Siripornadulsil, Sukon, & Siripornadulsil, 2014; Samocha et al., 2004; Shobharani, Padmaja, & Halami, 2015). Furthermore, fish and shrimp feed production processes require high temperatures and only spores from Bacillus species support this kind of conditions (Balca´zar

Bacillus Probiotic Enzymes

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Gras Antibiotics Fast growth rate Bacillus subtilis Spores

Aerobic Anaerobic Proteases Carbohydrases lipases

Fig. 8 Bacillus subtilis intrinsic properties.

et al., 2006; Valdez et al., 2014). B. subtilis enzymes also support extreme environmental conditions increasing animal capabilities to digest and assimilate nutrients from several origins including plants (Arellano-Carbajal & Olmos-Soto, 2002; Hamza, Fdhila, Zouiten, & Masmoudi, 2015; Ochoa-Solano, 2012; Ochoa-Solano & Olmos, 2006, Olmos et al., 2011, 2014). The capacity of B. subtilis to grow aerobically and anaerobically allows this bacterium to survive and grow adequately inside of animals and in aquaculture water systems (Fig. 8). All these properties give B. subtilis the potential capacity to become “one of the most perfect multifunctional probiotic bacteria” to formulate FFs for aquaculture (Olmos et al., 2014).

8. CONCLUSIONS Because forage fish species are considered to be keystones of their ecosystems, it is important to harvest them responsibly. Fish meal and fish oil have been and will continue to be vital ingredients in many kinds of aquaculture diets. Although supplies are likely to remain scarce, some sectors of aquaculture will be able to grow by complementing the marine ingredients

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with other resources. This will result in lowering inclusions of both fish meal and fish oil and will promote plant ingredients additionally, increasing digestive and diseases’ problems but lowering prices and improving sustainability of the seas. B. subtilis is “one of the most perfect multifunctional probiotic bacteria” that can help to solve aquaculture problems induced by vegetable ingredients’ inclusion. Thus, the development of new generation of safe, economic, and environmentally friendly diets will be based on this bacterium.

ACKNOWLEDGMENT The author would like to thank Alejandro Sanchez Gonzales and Rosalia Contreras Flores for editing tables and figures.

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CHAPTER THREE

Characterization and Applications of Marine Microbial Enzymes in Biotechnology and Probiotics for Animal Health T.H. Nguyen*,1, V.D. Nguyen†,1 *Faculty of Food Technology, Nha Trang University, Nha Trang, Vietnam † Institute of Biotechnology and Environment, Nha Trang University, Nha Trang, Vietnam 1 Corresponding authors: e-mail address: [email protected]; [email protected]

Contents 1. Introduction 2. Isolation, Purification, and Characterization of Marine Microbial Enzymes 2.1 Protease 2.2 Lipase 2.3 Polysaccharide-Degrading Enzymes 3. Marine Microbial Enzymes as Tools in Biotechnology 3.1 Molecular Biology Applications 3.2 Clinical and Other Health-Related Applications 4. Marine Microbial Enzymes as Tools in Probiotics to Benefit Human and Animal Health 5. Conclusions References

38 38 39 41 43 49 49 51 56 63 64

Abstract Marine microorganisms have been recognized as potential sources of novel enzymes because they are relatively more stable than the corresponding enzymes derived from plants and animals. Enzymes from marine microorganisms also differ from homologous enzymes in terrestrial microorganisms based on salinity, pressure, temperature, and lighting conditions. Marine microbial enzymes can be used in diverse industrial applications. This chapter will focus on the biotechnological applications of marine enzymes and also their use as a tool of marine probiotics to improve host digestion (food digestion, food absorption, and mucus utilization) and cleave molecular signals involved in quorum sensing in pathogens to control disease in aquaculture.

Advances in Food and Nutrition Research, Volume 80 ISSN 1043-4526 http://dx.doi.org/10.1016/bs.afnr.2016.11.007

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2017 Elsevier Inc. All rights reserved.

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1. INTRODUCTION Enzymes have been isolated and purified from microorganisms, plants, and animals, and their properties differ markedly depending on their respective sources. Microorganisms represent the most common source of enzymes because of their broad biochemical diversity, feasibility of mass culture, and ease of genetic manipulation (Niehaus, Bertoldo, K€ahler, & Antranikian, 1999; Shi, Song, & Zhang, 2013). Microorganisms are preferred to plants and animals as sources of enzymes because of economic and technical reasons (Gurung, Ray, Bose, & Rai, 2013; Vijayan, Swapna, Haridas, & Sabu, 2016). The oceans cover more than three quarters of the Earth’s surface and provide abundant resources for biotechnological research and development. Marine microorganisms have been recognized as potential sources of novel enzymes due to their better stability, activity, and tolerance to extreme conditions that most of the other proteins cannot withstand (Littlechild, 2015; Trincone, 2011). The complexity of the marine environment involving high salinity, high pressure, low temperature, and special lighting conditions may contribute to the significant differences between the enzymes generated by marine microorganisms and homologous enzymes from terrestrial microorganisms (Sana, 2013; Zhang & Kim, 2010). A variety of enzymes have been isolated from marine bacteria, actinomycetes, fungi, and other marine microorganisms. Marine microbial enzymes have been reported to have a wide range of potential applications such as in the dairy, food, detergents, textile, pharmaceutical, cosmetic, and biodiesel industries. They are also used in the synthesis of fine chemicals, agrochemicals, and new polymeric materials (Sana, 2013). Usually, enzymes have to be purified prior to their applications and the choice of purification method depends on the type of enzyme and microorganism which is used as a source of the enzyme. This chapter discusses the purification, characterization, and applications of marine microbial enzymes in biotechnology and probiotic technology.

2. ISOLATION, PURIFICATION, AND CHARACTERIZATION OF MARINE MICROBIAL ENZYMES A variety of enzymes with specific activities have been isolated from marine bacteria, fungi, and yeasts. In this section, the production, purification, and characterization of proteases, lipases, and polysaccharide-degrading enzymes from marine microorganisms are discussed.

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2.1 Protease Proteases represent 60% of total of the worldwide sales. They have wide applications in pharmaceutical, leather, laundry, food, and waste processing industries (Zhang & Kim, 2010). Two extracellular proteases were early reported from marine Pseudomonas strain 145-2 by ammonium sulfate fractionation following Sephadex gel chromatography from Makino, Koshikawa, Nishihara, Ichikawa, and Kondo (1981). In another early study by Qua, Simidu, and Taga (1981), a halophilic protease in culture fluids of a moderately halophilic marine Pseudomonas sp. (A-14) was purified by ammonium sulfate precipitation, DEAE-cellulose chromatography, and gel filtration through Sephadex G-200. The enzyme was purified to homogeneity, as judged from polyacrylamide gel electrophoresis. It had a molecular weight of 120 kDa. It showed optimal activity at 18% NaCl and pH 8.0. Mg2+, Co2+, and Ca2+ were found to act as activators, while Fe2+, Cu2+, and Hg2+ inactivated the enzyme activity. Qiu, Li, Dai, and Yuan (1991) selected 30 marine bacteria strains from the sea water, mud, fish, and other samples and screened them for proteolytic activity. Protease was best produced by the N1–35 strain. This strain produced protease that had significant advantages compared with the terrestrial ones. An alkaline protease with a relative molecular mass of 36 kDa was isolated from marine shipworm bacterial cultures (Griffin, Greene, & Cotta, 1992). The cultures were harvested after 10 days of growth by centrifugation. The supernatant obtained was ultracentrifuged. Then, the clear, membranous pellet obtained from ultracentrifugation was discarded. The supernatant was concentrated by ultrafiltration and then diafiltered with distilled water. The combined filtrate and diafiltrate were concentrated by ultrafiltration and then were precipitated with ammonium sulfate. The precipitate was resuspended in an aqueous NaC1 solution containing HEPES buffer. Aliquots of the solution were applied to an HPLC system fitted in series with a Bio-Sil SEC 250 column and a Bio-Sil SEC 125 column. Fractions containing protease activity were assayed for proteolytic activities. It was found that the enzyme was activated and stabilized by relatively high salt concentrations (>0.2 M). The specific activity of the enzyme was 65,840 proteolytic U/mg using azocasein as a substrate at the optimal temperature of 42°C, pH of 9.0, and salt concentration of 0.20 M. The basic residues, histidine, arginine, and lysine, represented 10% of the protein hydrolysate. The sequence obtained for the first 10 residues of the enzyme was Ile-Val-Tyr-Pro-Arg-Val/Tyr-Ala-GlymetSer. The activity of the purified enzyme was stable up to 40°C from pH 3.0 to 11.9, and from 0.1 to 3.5 M NaCl. These characteristics suggested a potential industrial application of the isolated protease. Patel, Dodia,

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Joshi, and Singh (2006) isolated and identified the haloalkaliphilic, Grampositive, aerobic, coccoid Bacillus pseudofirmus-Po2 by 16S rRNA sequencing analysis from the seawater sample. Screening and optimization for the production of alkaline protease were investigated using NaCl, nitrogen sources, and metal ions. It was found that Po2 could grow optimally at 10% NaCl (w/v) and pH 8.0. The protease production was salt dependent and optimum production required 15% NaCl (w/v) and pH 8.0. Among the organic nitrogen sources, optimum growth and protease production were supported by the combination of peptone and yeast extract. However, growth and protease production were highly suppressed by inorganic nitrogen sources used. Strong inhibition of enzyme production was observed at above 1% glucose (w/v). A yeast strain (Aureobasidium pullulans) showing the proteolytic activity was isolated from sea saltern of the China Yellow Sea (Chi, Ma, Wang, & Li, 2007). The optimum fermentation conditions for enzyme production were observed to be a medium containing 2.5 g soluble starch and 2.0 NaNO3; 100 mL seawater; initial medium, pH of 6.0; incubation temperature of 24.5°C; and incubation period of 30 h. Ma, Ni, Chi, Ma, and Gao (2007) purified an extracellular alkaline protease in the supernatant of cell culture of this yeast strain. Purification was achieved by ammonium sulfate fractionation, gel-filtration chromatography (Sephadex™ G-75), and anion-exchange chromatography (DEAE Sepharose Fast Flow). The molecular mass of the purified enzyme was estimated to be 32.0 kDa. The purified enzyme had a maximum activity at pH 9.0 and 45°C. The enzyme was activated by Cu2+ and Mn2+ and inactivated by Hg2+, Fe2+, Fe3+, Zn2+, and Co2+. Haddar et al. (2009) purified two alkaline serine proteases (BM1 and BM2) from Bacillus mojavensis A21, which was isolated from seawater. The molecular weights of BM1 and BM2 enzymes determined by SDS-PAGE were approximately 29 and 15.5 kDa, respectively. The optimum pH values of BM1 and BM2 proteases were found to be 8.0–10.0 and 10.0, respectively. Both enzymes exhibited maximal activity at 60°C, using casein as a substrate, and showed high stability toward nonionic surfactants. In addition, both of them showed excellent stability and compatibility with a wide range of commercial liquid and solid detergents. An extracellular alkaline protease produced by marine bacteria strain Pseudoalteromonas sp. 129-1 was purified by ammonium sulfate precipitation, anion-exchange chromatography, and gel filtration (Wu, Liu, Zhang, Li, & Sun, 2015). The molecular mass of the purified protease was estimated to be 35 kDa. The protease maintained considerable activity and stability at a wide temperature range of 10–60°C and pH range of 6.0–11.0.

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It had an optimal activity at the temperature of 50°C and pH of 8.0 and showed high stability in the presence of surfactants (SDS, Tween 80, and Triton X-100), oxidizing agent H2O2, and commercial detergents. Recently, a serine protease-producing marine bacterial strain PT-1 was isolated and identified as a family of Marinomonas arctica (Yoo & Park, 2016). Optimized culture conditions for the growth of the bacterium PT-1 and production of protease were determined to be pH 8.0 in the presence of 5% NaCl, at 37°C during 24 h of incubation. The molecular weight of the purified protease was estimated to be 63 kDa. The purified enzyme was completely inhibited by phenylmethylsulfonyl fluoride. Besides bacteria, various proteases have been purified and characterized from marine fungi (Budiarto, Mustopa, & Tarman, 2015; Indarmawan, Mustopa, Budiarto, & Tarman, 2016; Kamat, Rodrigues, & Naik, 2008) and from marine yeasts (Lario et al., 2015; Li, Peng, Wang, & Chi, 2010).

2.2 Lipase Lipases are classified as hydrolases that catalyze the breakdown of lipids with subsequent release of free fatty acids, diacylglycerols, monoglycerols, and glycerol. They are also efficient in various reactions such as esterification, transesterification, and aminolysis (Villeneuve, Muderhwa, Graille, & Haas, 2000). Microbial lipases are relatively stable and are capable of catalyzing a variety of reactions, being of importance for diverse industrial applications. The production of microbial lipases was first found from Penicillium oxalicum and Aspergillus flavus in 1935 by David. Later, four cold-adapted lipases were screened from Moraxella (Feller, Thiry, Arpigy, Mergeay, & Gerday, 1990). These Moraxella were obtained from the Antarctic seawater with the optimum growth temperature of 25°C, the maximum secretion of lipases was found to occur at lower temperature conditions, and the lowest secretion temperature can reach 3°C. A total of 427 yeast strains isolated from seawater, sediments, mud of salterns, guts of the marine fish, and marine algae were screened for lipase activity (Wang, Chi, Wang, Liu, & Li, 2007). Among them, nine yeast strains could produce lipase including Candida intermedia YA01a, Pichia guilliermondii N12c, Candida parapsilosis 3eA2, Lodderomyces elongisporus YF12c, Candida quercitrusa JHSb, Candia rugosa wl8, Yarrowia lipolytica N9a, Rhodotorula mucilaginosa L10-2, and A. pullulans HN2.3. The optimal pHs and temperatures of lipases produced by the isolated strains were between 6.0 and 8.5 and between 35 and 40°C, respectively. With the exception of lipase from A. pullulans HN2.3, which

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was extracellular, all lipases from the other yeast strains were cell bound. A novel extracellular phospholipase C was purified from a marine streptomycete, which was selected from approximately 400 marine bacteria by Mo, Kim, and Cho (2009). Its enzyme activity was optimal at pH 8.0 at 45°C, and it hydrolyzed only phosphatidylcholine. The enzyme activity was enhanced 300% by Na+ (200 mM), suggesting that the purified phospholipase C is a typical marine-type enzyme. Basheer et al. (2011) found that marine fungus Aspergillus awamori BTMFW032 isolated from seawater could produce extracellular lipase. Medium with soybean meal, 0.77% (w/v); (NH4)2SO4, 0.1 M; KH2PO4, 0.05 M; rice bran oil, 2% (v/v); CaCl2, 0.05 M; PEG 6000, 0.05% (w/v); NaCl, 1% (w/v); inoculums, 1% (v/v); pH 3.0; incubation temperature, 35°C; and incubation period, 5 days were identified as optimal submerged fermentation conditions for maximal lipase production. Partial purification by (NH4)2SO4 precipitation and ion exchange chromatography resulted in 33.7% final yield. The isolated lipase had a molecular mass of 90 kDa and an optimal activity at pH 7.0 and 40°C. A total of 44 marine sediments from 8 locations along the South East Coast of Bay of Bengal, India were screened for lipolytic fungal isolates by tributyrin agar clearing method and submerged fermentation (Bindiya & Ramana, 2012). It was found that marine fungus Aspergillus sydowii BTSS 1005 could produce higher extracellular lipase when compared with other isolates. The optimum medium composition under submerged fermentation for this strain was found to be sucrose, 2% (w/v); ammonium chloride, 3.5% (w/v); olive oil, 3% (v/v); and Tween 80, 0.2% (v/v). This research group did not purify and characterize the lipase. In the study of Smitha, Correya, and Philip (2014), lipase production of 181 marine fungi isolated from the continental slope sediments of Arabian Sea was investigated. Among the 181 cultures, 60.2% of fungi showed lipase activity. Thermostability is a desirable characteristic of lipase because it may allow the enzymatic reaction to be performed at higher temperature and would be helpful to increase substrate solubility and conversion rates, and to reduce the viscosity of reaction medium and the contamination of microorganism (Li & Zhang, 2005). Kiran, Lipton, Kennedy, Dobson, and Selvin (2014) purified and characterized a halotolerant thermostable lipase from the marine bacterium Oceanobacillus sp. PUMB02. The lipase was partially purified by ammonium sulfate precipitation and then was subjected to dialysis, butyl-Sepharose separation, anionexchange chromatography on DEAE cellulose, and gel-filtration chromatography on Sephadex G-75 column. The purified lipase showed optimal activity at 30°C and pH 8.0 and was stable at higher temperatures

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(50–70°C) and alkaline pH. The molecular mass of the purified lipase was estimated to be 31 kDa based on SDS-PAGE and MALDI-ToF fingerprint analysis. Yuan, Lan, Xin, Yang, and Wang (2015) reported a potential thermostable lipase (MAS1) from marine Streptomyces sp. strain W007, which was stable for 1-h incubation at temperature up to 60°C, suggesting that MAS1 was a thermostable lipase. In addition, the lipase MAS1 showed the high tolerance to organic solvents; it retained more than 90% of activity in methanol, alcohol, isopropanol, and acetone. These characteristics suggested that the lipase MAS1 was considered as a potential biocatalyst for industrial application. However, the production level of MAS1 was relatively low. Later, the same research group improved the expression level of MAS1 lipase in Pichia pastoris via chaperon coexpression and fermentation condition optimization (Lan, Qu, Yang, & Wang, 2016). It was found that the expression level of lipase MAS1 increased to 442 U/mL, which were about 44-fold to that of shaking flask cultivation (10 U/mL). Recently, several groups have reported the lipase production from various marine bacteria (Nerurkar, Joshi, & Adivarekar, 2015; Teymouri, Karkhane, Gilavand, Akhtari, & Marzban, 2016), marine fungi (Patnala, Kabilan, Gopalakrishnan, Rao, & Kumar, 2016; Sadati, Barghi, & Abbasi, 2015), and marine yeasts (Louhasakul, Cheirsilp, & Prasertsan, 2016).

2.3 Polysaccharide-Degrading Enzymes According to Zhang and Kim (2010), the total production of chitin in the whole marine biocycle is at least 2.3 million metric tons per year. Several marine bacteria, including Aspergillus, Penicillium, Rhizopus, Myxobacter, Sporocytophaga, Bacillus, Enterobacter, Klebsiella, Pseudomonas, Serratia, Chromobacterium, Clostridium, Flavobacterium, Arthrobacter, and Streptomyces, have been reported to produce chitinase or chitosanase (Fukamizo, 2000; Monzingo, Marcotte, Hart, & Robertus, 1996; Xia, Liu, & Liu, 2008). A total of 48 bacterial strains capable of utilizing chitin as a sole source of nutrients were isolated from river and marine waters in Tokushima, Japan (Osama & Koga, 1995). These bacteria were identified as Vibrio fluvialis, Vibrio parahaemolyticus, Vibrio mimicus, Vibrio alginolyticus, Listonella anguillarum, and Aeromonas hydrophila. All strains were able to produce chitinase. Suresh and Chandrasekaran (1999) reported chitinase production from the marine strains of Beauveria bassiana (Bals.) Vuill. by solid-state fermentation. The strain was strongly alkalophilic and produced maximum chitinase at pH 9.2. The purification of chitinases and chitosanases from

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microbial sources in most cases has involved classical enzyme purification methods. These methods involve removal of microbial biomass from the culture broth, followed by selective precipitation/concentration by (NH4)2SO4 or solvents or polyethylene glycol (Thadathil & Velappan, 2014). The concentrated chitosanase is further subjected to chromatography, commonly gel filtration, ion exchange, affinity techniques, etc. Chitinase-producing bacterial strain, Micrococcus sp. AG84, isolated from marine sediments grew maximally in a shake flask and produced chitinase at 35°C and pH 8.0 (Annamalai, Giji, Arumugam, & Balasubramanian, 2010). For purification of the enzyme from AG84, the cell-free supernatant was first precipitated with ammonium sulfate at 60% saturation. The precipitates were collected by centrifugation and dialyzed against Tris–HCl buffer. The partially purified enzyme was obtained by a DEAE-cellulose column equilibrated with Tris–HCl buffer. The partial purified chitinase was found to be the maximum at 45°C and pH 8.0. It was noted that the purified enzyme was 100% stable even at 60°C and pH 11.0. The presence of Fe2+, Ca2+, and Ni2+stimulated chitinase activity, whereas the activity was inhibited by EDTA. The molecular weight of the purified chitinase was 33 kDa. Velmurugan, Duraisamy, Hoon, Jung, and Yang (2011) reported a novel low-temperature chitinase from the marine fungus Plectosphaerella sp. strain MF-1. The strain MF-1 was isolated from sea shells and chitinase production by this strain was detected by inoculating the fungus into M9 medium containing 0.5% colloidal chitin at 10°C. The crude chitinase solution was purified by ammonium sulfate fractionation, cellulose DEAE anion exchange, and Sephadex gel-filtration chromatography. The purified chitinase had the maximal activity at 37°C. Interestingly, the difference in chitinase activities at 10°C and 37°C was less than 0.01 U/mL, indicating that chitinase was active at low temperature. The optimal pH for the low-temperature active chitinase was 3.0–4.0. The Km was 0.03 mM and Vmax was 0.095, using p-nitrophenyl N-acetyl-β-D-glucosaminide as a substrate. Several metal ions including Ag+, Hg2+, and Pb2+ significantly inhibited enzyme activity, whereas Mg2+ and Fe2+ had minimal inhibition. The molecular mass of purified chitinase was determined to be 67 kDa by SDS-PAGE. The N-terminal amino acid sequence was determined to be “DNISQTGEHARYXPMVWFIKL.” A variety of chitinase genes have already been cloned from marine bacteria and fungi. For instance, Suolow and Jones (1988) inserted two chitinase genes (ChiA, ChiB) into Escherichia coli, and subsequently, these genes were transferred into Pseudomonas; finally, they acquired four high-yielding chitinase strains. Screening of

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extracellular chitosanase from bacterial isolates associated with marine sponges was reported by Chasanah, Zilda, and Uria (2009). Accordingly, out of 100 bacterial isolates, 40 isolates were capable of forming clearing zones on the chitin media and 1 isolate, 34-b, produced the highest chitinolytic index. The enzyme was produced on chitin liquid medium at 37°C in a shaking water bath for a 5-day cultivation. Crude enzymes were prepared by cell-free supernatant and concentrated through 70% ammonium sulfate precipitation followed by dialysis. The enzymes worked best at pH and temperature of 6–7 and 60°C, respectively. The half-life (T1/2) for chitosanase activity was 500.2 min (at 37°C) and 55.12 min (at 50°C), indicating that the enzyme is quite stable at that temperature. However, around 80% of the original activity was lost at 60°C after 15 min of incubation. An extracellular chitosanase was purified from marine bacterium Bacillus subtilis CH2 (Chulhong et al., 2011). The strain was isolated from the intestine of Sebastiscus marmoratus (scorpion fish). The chitosanase of B. subtilis CH2 was best induced by fructose and was not induced with chitosan, which was different from other found chitosanases. The strain was incubated in LB broth, and the chitosanase secreted into the medium was first concentrated with ammonium sulfate precipitation and then purified by gel permeation chromatography. The molecular mass of the purified chitosanase was determined as 29 kDa. The optimum pH and the temperature of the purified chitosanase were 5.5 and 60°C, respectively. The specific activity of the purified chitosanase was 161 U/mg. Recently, a novel chitinase gene (PbChi70) from a marine bacterium Paenicibacillus barengoltzii has been cloned and functionally expressed in E. coli (Yang et al., 2016). The molecular mass of the purified enzyme was estimated to be 70.0 kDa. PbChi70 displayed maximal activity at pH 5.5 and 55°C. It exhibited strict substrate specificity for colloidal chitin, glycol chitin, powdery chitin, and N-acetyl chitooligosaccharides. A thermostable and alkali tolerant from marine bacterial Bacillus pumilus JUBCH08 was recently reported by Bhattacharya, Das, Samadder, and Rajan (2016). It showed a maximum activity at pH 8.0 and 70°C for 1 h. The molecular weight of chitinase was found to be 64 kDa, and its activity was improved in the presence of Mg2+, Co2+, Ca2+, and Mn2+. Very recently, Fu, Yan, Wang, Yang, and Jiang (2016) reported a novel acidic chitinase (PbChi67) from the marine bacterium P. barengoltzii CAU904. The molecular mass of the enzyme was 67.0 kDa by SDS-PAGE and 67.9 kDa by gel filtration, respectively. PbChi67 was most active at pH 3.5 and was stable within pH 3.0–9.0 and up to 55°C. It hydrolyzed colloidal chitin to yield N-acetyl chitooligosaccharides.

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Agar is a highly heterogeneous polysaccharide extracted from the cell wall of a group of red algae (Rhodophyceae), including Gelidium and Gracilaria. Agar is formed by a mixture of two polysaccharides named agarose and agaropectin (Araki, 1937). The main structure of agarose is composed of repetitive units of β-D-galactose and 3,6-anhydro-α-L-galactose (3,6-AG), with few variations, and a low content of sulfate esters. Agar oligosaccharides have a wide range of applications in the food industry; it can be used for beverages, bread, and some low-calorie food production (Zhang & Kim, 2010). Agarases catalyze the hydrolysis of agar. They are classified into α-agarase (E.C. 3.2.1.158) and β-agarase (E.C. 3.2.1.81) according to the cleavage pattern. The basic structure of agar is composed of repetitive units of β-D-galactose and 3,6-anhydro-α-L-galactose (Araki, 1937). The sources, isolation, characterization, and application of agarase have been reviewed by Fu and Kim (2010). Agarases are usually purified by a combination of several techniques, including ammonium sulfate precipitation followed by gel filtration and anion-exchange chromatography. In some cases, ammonium sulfate fractionation may be omitted (Fu, Lin, & Kim, 2008) or replaced by acetone precipitation (Ha et al., 1997) in the purification procedure. Anion exchangers were used in the purification procedure for capturing agarase, indicating that agarases are proteins with pI values lower than 7 (Fu & Kim, 2010). Agarases are obtained mainly from marine bacteria. To date, several microorganisms isolated from seawater and marine sediments have been reported to produce agarases, including Vibrio (Doi, Chinen, Fukuda, & Usuda, 2016; Zhang & Sun, 2007), Pseudomonas (Hsu et al., 2015; Ryu et al., 2001), Alteromonas (Chi et al., 2014; Wang, Mou, Jiang, & Guan, 2006), Microbulbifer (Jonnadula & Ghadi, 2011; Vijayaraghavan & Rajendran, 2012), Thalassomonas (Jean, Shieh, & Liu, 2006; Liang, Chen, Chen, Chiu, & Liaw, 2014), Salegentibacter (Nedashkovskaya et al., 2006), Zobellia (Hehemann et al., 2012; Jam et al., 2005), Agarivorans (Fu, Pan, Lin, & Kim, 2009; Lin et al., 2012), Pseudoalteromonas (Chi et al., 2015; Oh, Jung, & Lee, 2011), and Paenibacillus (Song et al., 2014). Sugano, Terada, Arita, and Noma (1993) reported early the purification and characterization of an endo-type beta-agarase (agarase 0107) from a marine bacterium Vibrio sp. JT0107. This enzyme was purified by a combination of ammonium sulfate precipitation and successive rounds of anion-exchange column chromatography. The purified agarase had a molecular mass of 107 kDa and an isoelectric of 6.3. The optimum pH and the temperature for enzyme activity were pH 8.0 and 30°C, respectively. With respect to amino acid composition, aspartic acid is the most

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abundant, followed by alanine and glutamic acid. Hu, Lin, Xu, Zhong, and Liu (2009) reported production and purification of different agarases from a marine agarolytic bacterium Agarivorans sp. HZ105. The optimum conditions required for maximum production were found to be pH of 8.0, 2.5% NaCl, and cultivation period of 16 h. A quick and simple method (eclaim protein from the polyacrylamide gel after PAGE) was successfully applied to separate two agarases of strain HZ105. Agarases were purified to homogeneity directly from bands in the PAGE gel. The molecular mass of the two purified agarases was estimated to be 58 and 54 kDa, respectively. Based on the MS spectra result, all agarases of strain HZ105 are beta-agarase and belong to the family 50 of glycosyl hydrolases. Another extracellular beta-agarase (AgaA34) was purified from an isolated marine bacterium, Agarivorans albus YKW-34 from the gut of a turban shell (Fu et al., 2008). AgaA34 was purified to homogeneity by ion exchange and gel-filtration chromatographies with a recovery of 30% and a fold of 10. AgaA34 was composed of a single polypeptide chain had a molecular mass of 50 kDa. N-terminal amino acid sequencing revealed a sequence of ASLVTSFEEA, indicating that the purified agarase belongs to glycoside hydrolase family 50. The pH and the temperature optimum of AgaA34 were pH 8.0 and 40°C, respectively. It was stable over pH 6.0–11.0 and at the temperature up to 50° C. Metal ions were not required for its activity, while reducing reagents (beta-Me and dithiothreitol, DTT) increased its activity by 30%. Various other Agarivorans species were also reported to have the ability to produce agarase and catalyze the hydrolysis of agar (Liu, Mao, Du, Mu, & Wei, 2014; Long, Yu, & Xu, 2010). Notably, an agarase AgaA obtained from the marine bacterium Agarivorans sp. LQ48 could retain more than 95% its activity after incubation at pH 3.0–11.0 for 1 h, a characteristic much different from other reported agarases (Long et al., 2010). Recently, Zhang et al. (2016) presented the complete genome sequence of A. albus and Agarivorans gilvus WH0801. The genome included two β-agarase genes and three glycosyl hydrolase genes. β-Agarase produced by Paenibacillus sp. WL (agarase WL) was purified using a combination of ammonium sulfate precipitation, DEAE-ion exchange, and gel-filtration chromatography (Mei et al., 2014). The molecular mass of the purified agarase was approximately 30 kDa. The agarase was stable at the temperature below 50°C and the favorable agar-hydrolysis activity was at 40°C. The agarase was active in the range of pH 5.0–8.0, and the optimal agar-hydrolysis pH value was 6.0. Various metal ions normally found in seawater (Na+, K+, Ca2+, Mg2+, and Al3+) could activate agarase WL. Recently, a putative agarase gene

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(ragaH92), an endo-type beta-agarase, was identified in an agarolytic marine bacterium, Pseudoalteromonas sp. H9 (Chi et al., 2015). The molecular mass of rAgaH92 was 51 kDa and the optimum pH and the temperature for rAgaH92 activity were 6.0 and 45°C, respectively. The agarase activity of rAgaH92 was enhanced by Fe2+ and strongly inhibited EDTA. Another endo-type β-1,4 agarase was also recently obtained from the isolated marine bacterium Pseudomonas vesicularis MA103 (Hsu et al., 2015). The extracellular production of recombinant P. vesicularis AgaA was showed to have agarase activity higher than that of cytosolic AgaA. A novel gene (Aga4436), encoding a potential agarase of 456 amino acids, was recently reported from the genome of deep-sea bacterium Flammeovirga sp. OC4. Aga4436 (Chen, Hou, Jin, Zeng, & Lin, 2016). Recombinant Aga4436 was active over a broad range of pHs (5.0–10.0) and temperatures (30–80°C) and retained more than 90%, 80%, and 35% of its maximum activity after incubation at 30°C, 40°C, and 50°C for 144 h, respectively. Additionally, Aga4436 displayed a remarkable tolerance to acid and alkaline environments, as it retained more than 70% of its maximum activity at a wide range of pHs from 3.0 to 10.0 after incubation for 60 min. This suggested its potential application in the food and nutraceutical industries. Alginate is an abundant polysaccharide found within the cell wall of brown, which consists of β-D-mannuronate (M) and α-L-guluronate (G) as monomeric units. Alginate has a wide range of applications in food and pharmaceutical industries. Alginate lyases, characterized as either mannuronate or guluronate lyases, are a complex copolymer of α-L-guluronate and its C5 epimer β-D-mannuronate. It can degrade alginate by β-elimination of glycosidic bonds and produce unsaturated oligosaccharides with double bonds at the nonreducing end. Alginate lyases can be grouped into three types based on their molecular masses: small (25–30 kDa), medium-sized (around 40 kDa), and large lyases (>60 kDa) (Zhu & Yin, 2015). Alginate lyases have been isolated from a wide range of organisms, including marine and terrestrial bacteria, marine mollusks, and algae. In recent years, the marine microbial alginate lyases have been greatly studied. A high-alkaline, salt-activated alginate lyase (A1m) produced by Agarivorans sp. JAM-A1m from a deep-sea sediment off Cape Nomamisaki on Kyushu Island (Japan) has been reported by Kobayashi, Uchimura, Miyazaki, Nogi, and Horikoshi (2009). The purified enzyme had a molecular mass of approximately 31 kDa by SDS-PAGE. The optimal pH and the temperature for enzyme activity were found to be around 10.0 and 30°C, respectively, and the activity was increased by adding 0.2 M NaCl. Several other

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salt-activated alginate lyases have been reported from marine bacteria (Kitamikado, Tseng, Yamaguchi, & Nakamura, 1992; Uchimura, Miyazaki, Nogi, Kobayashi, & Horikoshi, 2010; Xiao, Feng, Yang, Lu, & Yu, 2006). Recently, Chen, Dong, et al. (2016) characterized an alginate lyase, AlyPM, from Pseudoalteromonas sp. SM0524. The optimal pH and the temperature for AlyPM activity were 8.5 and 30°C, respectively. Thermal and salt stability studies reveal that AlyPM is a cold-adapted and saltactivated enzyme. Some cold-adapted alginate lyases have also been found to be secreted by bacteria from the Arctic (Dong et al., 2012) and other sea (Li, Yang, Zhang, Yu, & Han, 2015; Xiao et al., 2006). Very recently, Yagi, Fujise, Itabashi, and Ohshiro (2016) purified a novel thermostable and salttolerant alginate lyase (AlgC-PL7) from the halophilic Gram-negative bacterium Cobetia sp. NAP1, which was collected from the brown algae Padina arborescens Holmes. The bacterial cells were cultured in alginate medium at 20°C with shaking. Combination of centrifugation and heat treatment was applied to prepare crude enzyme. The crude enzyme was further dialyzed in Tris–HCl and loaded onto a Superdex 75 column to obtain AlgC-PL7. The purified enzyme was found to have a molecular weight of about 35 kDa according to SDS-PAGE and MALDI-FOF-MS results. AlgC-PL7 displayed its highest activity at pH 8.0 and relatively high temperature of 45°C. It is noted that the AlgC-PL7 could retain its activity in the temperature ranging from 30°C to 70°C.

3. MARINE MICROBIAL ENZYMES AS TOOLS IN BIOTECHNOLOGY Marine bacteria have to adapt to diverse environmental parameters such as high salt concentration, extreme temperature, acidic and alkaline pH, extreme barometric pressure, and low nutrient availability. Thus, marine microbial enzymes attract special interest due to their better stability, activity, and tolerance to extreme conditions that most of the other enzymes cannot withstand (Sana, 2013). Many marine microbial enzymes have been and are being used in biotechnology. This section will review potential applications of marine bacterial enzymes in molecular biology, and in clinical and other health-related fields.

3.1 Molecular Biology Applications Marine microorganisms are expected to have unique enzyme systems since they have to adapt to extreme marine environment condition such as high or

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low temperature, alkaline or acidic water, high pressure, and limited substrate in the deep-sea water. The nucleic acid enzymes, such as DNA polymerase, ligase, and restriction endonuclease, have important applications in molecular biology research. A thermostable DNA polymerase was purified from marine archaebacterium (Pyrococcus furiosus), which has sequence homology with α-like DNA polymerases (Mathur, Adams, Callen, & Cline, 1991). The DNA polymerase from P. furiosus has the lowest error rate of any known polymerase in polymerase chain reaction (PCR) amplification (Lu & Erickson, 1997). Thus, this polymerase can be applied in high-fidelity PCR experiments. A novel thermostable nonspecific nuclease from Thermophilic bacteriophage GBSV1 was purified from thermophilic bacteriophage GBSV1 isolated from an offshore hot spring in Xiamen of China. The enzyme can degrade various nucleic acids, including RNA, single-stranded DNA, and double-stranded DNA (Song & Zhang, 2008). Recently, Kwon et al. (2016) cloned, expressed, and characterized Tba5 DNA polymerase from Thermococcus barophilus Ch5, which was isolated from a deep-sea hydrothermal field of the Mid-Atlantic Ridge. The enzyme showed 30 ! 50 exonuclease activity. PCR amplification using Tba5 DNA polymerase enables high-yield for 1- to 6-kb target DNA products, while 8- to 10-kb target DNA products were amplified at low or inefficient levels. DNA ligase catalyzes the sealing of 50 -phosphate and 30 -hydroxyl termini at single-strand breaks in double-stranded DNA. It plays an important role in DNA replication and DNA repair pathways (Timson, Singleton, & Wigley, 2000). Pfu DNA ligase was isolated from the hyperthermophilic marine archaeon P. furiosus. This enzyme catalyzes the synthesis of adenosine 50 -tetraphosphate and dinucleoside polyphosphates by displacement of the adenosine 50 -monophosphate from the Pfu DNA ligase–AMP complex with tripolyphosphates, nucleoside triphosphates, or nucleoside diphosphates (G€ unther et al., 2002). Williamson, Rothweiler, and Schrøder Leiros (2014) reported the biochemical and structural characterization of a recombinantly produced ATP-dependent DNA ligase from the marine bacterium Psychromonas sp. strain SP041. This enzyme has intrinsic ATP-dependent ligase activity for variety of double-stranded DNA substrates and has similar kinetic constants to other small ATP-dependent DNA ligases from viruses and bacteria. Restriction endonucleases, which recognize a specific base sequence on DNA molecules and cleave the DNA molecules within or near the base sequence, are widely used for gene manipulation and gene analysis and are recovered from procaryotic cells (Williams, 2003). A novel restriction enzyme DmaI was isolated from the marine bacterium Deleya marina

Characterization and Applications of Marine Microbial Enzymes

51

IAM 14114 (Mizuno, Suzuki, Yamada, Akagawa, & Yamasato, 1990). This enzyme was stable up to 55°C and between pH 7 and 9. The purified enzyme recognized the palindromic hexanucleotide DNA sequence 5ʹ-CAGCTG-3ʹ, cut between G and C, and produced a flush end. Several other restriction endonucleases, including AspMD1, DmaI, DpaI, AgeI, HjaI, Hac1, Hsa1, and Hag1, were recently isolated from marine bacteria and some of them are already commercially available (Muffler, Sana, Mukherjee, & Ulber, 2015). Agarases are brilliant biological tools for DNA recovery from agarose gel after electrophoresis. Several agarases, isolated from marine bacteria, have been used as a tool to recover DNA from agarose gel (Sugano, Matsumoto, Kodama, & Noma, 1993; Zhang & Sun, 2007).

3.2 Clinical and Other Health-Related Applications Clinical and medicinal applications of enzymes have been widely developed. As mentioned previously, microorganisms living in the sea must be able to survive and grow in the water environment with low nutrition, high salinity, and high pressure. These characteristics made marine microorganisms able to produce the most potent source of antibiotics, bioactive enzymes, and other bioactive secondary metabolites. Many studies have shown that alkaline proteinase in the intestine of marine animals plays an important role in the digestion of protein in the feed. Angiotensin-I-converting enzyme (ACE) plays an important physiological role in the regulation of blood pressure via conversion of angiotensin I to angiotensin II. The peptides with ACE inhibitory activity have been reported to be used in the treatment of hypertension (He, Liu, & Ma, 2013). Zhao et al. (2008) reported the collagenolytic activity of deseasin MCP-01 secreted by a cold-adapted bacterium Pseudoalteromonas sp. SM9913 from the deep-sea sediment. MCP-01 had a broad substrate specificity to various type collagens from terrestrial and marine animals. Moreover, MCP-01 had various but specific cleavage sites on insoluble collagen, and thus insoluble collagen fiber was digested into dissolved small peptides. This suggested that MCP-01 would be used to produce collagen peptides with ACE inhibitory activity from fish processing by-products. An extracellular alkaline protease was purified from the marine yeast A. pullulans for bioactive peptide production from different sources (Ma et al., 2007). The enzymatic hydrolysis of shrimp protein, spirulina (Arthrospira platensis) protein, proteins of marine yeast strains N3C (Y. lipolytica) and YA03a (Hanseniaspora uvarum), milk protein, and casein with the purified extracellular alkaline protease showed to produce peptides

52

T.H. Nguyen and V.D. Nguyen

able to inhibit the ACE. The gene from the marine yeast A. pullulans HN2-3 was cloned into a surface display vector pINA1317-YlCWP110 and expressed in cells of Y. lipolytica as an extracellular protein. It was found that the overexpressed protease on Y. lipolytica surface could produce bioactive peptides with ACE inhibitory activity and antioxidant activity. An important application of enzymes is the preparation of bioactive compounds. Agarases are applied in the production of oligosaccharides from agar. Oligosaccharides from agar have many biological properties such as hepatoprotective potential (Chen, Yan, Zhu, & Lin, 2006), antioxidation (Wang, Hu, Nie, Yu, & Xie, 2016), antiinflammatory activity (Higashimura et al., 2013), and antitumor-promoting activity (Enoki et al., 2012), and therefore, they have potential applications in the food, cosmetic, and medical industries. Oh et al. (2010) reported that oligoagrosaccharides produced by β-agarase from the marine bacterium Pseudoalteromonas sp. AG4 showed whitening and antioxidant activities with no cytotoxicity. The authors suggested that the hydrolysis products could have potential use in cosmetic and medical industries. Chitooligomers (COS), the degraded compounds of chitosan, have been the subject of increased attention in terms of their pharmaceutical and medicinal applications, due to their nontoxic and high solubility properties as well as their positive physiological effects (Lodhi et al., 2014). Enzymatic depolymerizing of chitosan is very useful and a more environmentally friendly process for producing COS. Liang et al. (2016) have recently reported an endo-type chitosanase (CS038) from Bacillus mycoides TKU038 using squid pen powder. The chitosan oligosaccharides obtained from the hydrolysis of chitosan by CS038 exhibited 2,2-diphenyl-1-picrylhydrazyl (DPPH) radical scavenging and antiinflammatory capabilities. This suggested that CS038 has a potential application in COS production as a medical prebiotic. Some enzymes from marine microorganisms showed excellent anticancer and antimicrobial activities. L-Asparaginase plays a vital role in medical application, particularly in treatment of acute lymphoblastic leukemia as an effective antitumor agent (Batool, Makky, Jalal, & Yusoff, 2016). Twenty-one fungal strains were isolated from marine environment of the Red Sea coasts of Egypt by Faraga, Hassanb, Beltagy, and El-Shenawy (2015). Among them, Aspergillus terreus yielded the highest L-asparaginase specific activity. In the study of Nayak, Porob, Fernandes, Meena, and Ramaiah (2014), 71 marine bacterial DNA extracts were PCR screened for the gene encoding L-asparaginase (ansA). Over 62%

Characterization and Applications of Marine Microbial Enzymes

53

(44) of them were positive for ansA gene. The positive cultures were from genera Bacillus and Staphylococcus. Recently, Shakambari et al. (2016) reported the purification, characterization, and biological application of L-asparaginases from a marine isolate Bacillus tequilensis PV9W. The purified L-asparaginase had effective acrylamide degradation activity (6 IU/mL) and cytotoxicity on HeLa cells with relatively low IC50 value of 0.036 IU/mL. G2 arrest is known to attribute to deprivation of L-asparagine, an essential amino acid for protein synthesis. The L-asparaginase-treated cells showed a significant higher percentage of G2 phase (32%) compared to untreated control cells (3%) (Fig. 1). Hemolysis has been reported to be the major drawbacks of many drugs used in therapeutics for which a wide number of side effects of toxicity to blood cells are reported. The L-asparaginase purified from B. tequilensis PV9W was found to have no toxic effect on the erythrocytes even up to 6 IU. This suggested that L-asparaginase from B. tequilensis PV9W could be a potential anticancer candidate agent with the least amount of side effects. L-Glutaminase, an amidohydrolase, is gaining importance on account of its potential anticancer activity. The L-glutaminase production from a novel marine isolate Vibrio azureus strain JK-79 was reported by Kiruthika and Saraswathy (2013). L-Glutaminase produced by Alcaligenes faecalis KLU102, isolated from the marine realm (Bay of Bengal), showed significant cytotoxicity against HeLa, with an IC50 value of 12.5 μg/mL (Pandian, Deepak, Sivasubramaniam, Nellaiah, & Sundar, 2014). Chitinases have been receiving an increased attention due to their role in the biocontrol of fungal phytopathogens. Han, Yang, Zhang, Miao, and Li (2009) reported the antifungal activity of chitinase from the marine Streptomyces sp. DA11 isolated from a sponge from the South China Sea. The purified chitinase produced by marine-derived A. terreus inhibited the growth of various microbial species including Aspergillus niger, Aspergillus oryzae, Penicillium oxysporum, Rhizoctonia solani, Candida albicans, and Fusarium solani. The purified chitinase produced by marine-derived A. terreus inhibited the growth of several pathogenic bacteria such as Staphylococcus aureus, Salmonella typhi, and Pseudomonas aeruginosa (Faraga, Abd-Elnabey, Ibrahim, & El-Shenawy, 2016). Recently, an extracellular protease isolated from the marine fungus Xylaria psidii was reported to have antibacterial activity against B. subtilis and S. aureus (Indarmawan et al., 2016). These findings suggest that marine microbial enzymes could be used for biocontrol of pathogenic fungi and some pathogenic bacteria, and also useful in the treatment of infections caused by bacteria. In medicine, collagenases are used in wound healing,

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T.H. Nguyen and V.D. Nguyen

120

A

Cell survival (%)

100 80 60 40 20 0

0.00 0.01 0.01 0.03 0.05 0.10 0.20 0.30 0.40 Specimen_001-Con

B 80 70 60 Count

50 40 30

G1:80% S:17% G2:3%

20

i

10 0 50

100

150

200

PI-A Specimen_001-Pt

250 (× 1.000)

Count

150

100

G1:58 S:10 G2:32

50

ii 0 50

100

Cell cycle analysis

150 PI-A

200

250 (× 1.000)

Fig. 1 (A) Cytotoxic effect on HeLa cell lines of L-asparaginase purified from Bacillus tequilensis PV9W. The cells were treated in triplicates with each concentration for 24 h, and the cell viability was determined by the MTT assay. Error bars indicate SD of mean of three independent repeats in triplicates (n ¼ 9). (B) The effect of L-asparaginase purified from Bacillus tequilensis PV9W on the cell cycle analysis in HeLa cell line using flow cytometry (i) control and (ii) L-asparaginase treated. Reproduced by permission of the Royal Society of Chemistry (Shakambari et al., 2016).

Characterization and Applications of Marine Microbial Enzymes

55

treatment of diabetes, and enzymatic debridement. Marine microorganisms have recently emerged as excellent sources of collagenase (Abdel-Fattah, 2013; Mukherjee, Webster, & Llewellyn, 2009; Ran et al., 2013, 2014). Oxidative/nitrosative stress is the result of a disequilibrium in oxidant/ antioxidant, which reveals from continuous increase of reactive oxygen (ROS) and reactive nitrogen species (RNS) production (Maes, Galecki, Chang, & Berk, 2011). Antioxidant enzymes, including catalase (CAT), ascorbate peroxidase, peroxidase (POD), superoxide dismutase (SOD), and glutathione peroxidase (GPx), play a major role in reducing ROS/ RNS levels (Lei et al., 2016). About 90% of the worldwide ocean water is in the 0–3°C range where the solubility of oxygen is increased. Consequently, marine aerobic microorganisms are adapted for protection against ROS with their imperative antioxidant enzymes. Several antioxidant enzymes have been found in marine microorganisms. CAT genes have been identified in the complete genomes of marine cold-adapted bacteria Desulfotalea psychrophila, Colwellia psychrerythraea, and Pseudoalteromonas haloplanktis (Wang et al., 2013). Fu, Wang, Hao, Zhu, and Sun (2014) reported a high activity CAT from marine bacterium Acinetobacter sp. YS0810. The CAT showed high alkali stability and thermostability, demonstrating its good application in medical field. POD has been used as an important component of reagents for clinical diagnosis (Regalodo, Garcia-Almandarez, & Duarte-Vazquez, 2004). A total of 34 enzymes with biotechnological potential were screened in 374 isolates of marine bacteria, and POD was found in almost isolates (M€ uhling et al., 2013). SOD plays an important role in the protection of the cells against the harmful environment of free radicals. It has physiological importance in humans and other animals for medical treatment (Sheng et al., 2014). An SOD gene was classified as iron-containing superoxide dismutases (rPsSOD), which was purified from Antarctic microorganisms Pseudoalteromonas sp. ANT506 (Wang, Wang, et al., 2016). The purified rPsSOD exhibited some special catalytic properties, such as high-enzymatic activity at low temperature, well stability at pH, high-salt tolerance concentrations, and stress ability to some metal ions; this suggested its important roles in protecting against oxidative stress induced by extreme conditions. GPx catalyzes the reduction of oxidized glutathione using NADPH as the substrate to produce reduced glutathione (GSH), which is an important antioxidant molecule that helps maintain the proper reducing environment of the cell. Recently, Ji, Barnwell, and Grunden (2015) characterized a recombinant glutathione reductase from the psychrophilic Antarctic bacterium C. psychrerythraea. In vivo oxidative stress studies

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T.H. Nguyen and V.D. Nguyen

indicated that the C. psychrerythraea glutathione reductase was able to complement an E. coli glutathione reductase deletion strain. Enzymes have been used as catalysts in many pharmaceutical synthesis processes. Marine enzymes are of special interest for their unique catalytic properties, novel stereochemical properties, and solvent stability (Sana, 2013). Epoxide hydrolases have received much attention because they are cofactor-independent enzymes that are “easy to use” for catalyzing the hydrolysis of racemic epoxides to yield highly enantiopure epoxides and vicinal diols. Epoxides and vicinal diols are valuable chiral building blocks for the preparation of biologically active molecules, such as agrochemicals, drugs, and other pharmaceutical compounds (Kotik, 2012). Several marine microbial epoxide hydrolases have been reported to have potential enantioconvergent applications for the synthesis of drug intermediates with specific stereochemistry (Kang, Woo, Kang, Hwang, & Kim, 2008; Saini et al., 2015; Woo et al., 2010; Woo, Kang, Kwon, Nyun, & Lee, 2015). Esterases catalyze the hydrolysis and formation of short-chain fatty acid esters. They are used as stereo-specific biocatalysis in pharmaceutical production (Oda, Fukami, Yokoi, & Nakajima, 2015). The esterase from the marine isolate Y. lipolytica CL180 was overexpressed in E. coli, and the recombinant enzyme was used to produce levofloxacin by preferential hydrolysis of S-enantiomer from a racemic mixture of ofloxacin ester (Kim et al., 2007). In recent years, many other esterases have been isolated and characterized from marine bacteria (Jiang et al., 2012; Jiang, Zhang, Gao, & Hu, 2016; Lee, 2016; Lee, Heo, Lee, & Choi, 2014; Wu, Zhang, et al., 2015).

4. MARINE MICROBIAL ENZYMES AS TOOLS IN PROBIOTICS TO BENEFIT HUMAN AND ANIMAL HEALTH Gut microbiota of humans and animals consists of a complex set of microorganisms that live in the digestive tracts. Many of the mare beneficial bacteria that utilize hard-to-digest foods, produce nutrients and energy, and protect the hosts from foodborne pathogens. Pathogens, or harmful bacteria, also reside in the digestive tracts; they release toxins and are increasingly associated with a series of diseases. Therefore, a balance of beneficial and harmful bacteria contributes to human and animal health (Nguyen, 2016; Nguyen & Nguyen, 2016). In fact, many types of bacteria are competing in the gut, and the winners have potential to cause health problems in the host (Nguyen, 2016; Nguyen, Pham, Nguyen, Nguyen, & Hoj, 2014;

Characterization and Applications of Marine Microbial Enzymes

57

Pham, Ho, & Nguyen, 2014). The term “probiotics” was introduced by Parker (1974) to describe organisms and substances that contribute to intestinal microbial balance. Probiotics are vital culture of bacteria and fungi that, when introduced through feed, have a positive effect on health. According to FAO/WHO, probiotics are defined as “live microorganisms which when consumed in adequate amounts confer a health benefit on the host.” With regard to enzyme approach, enzymes may act as tools of probiotics to benefit human and animal health through two pathways: improving host digestion (food digestion, food absorption, and mucus utilization) and degrading molecular signals involved in quorum sensing (QS) in pathogens. Digestive enzymes from these probiotics including microorganisms of marine origin have been reviewed (Nguyen, 2016; Nguyen, Le, & Trang, 2013), indicating that probiotics might have a beneficial effect in the digestive process of human and animal hosts through degrading protein, chitin, cellulose, lipid, starch, and phytate. Thus, this chapter will focus on the second pathway involved in the potential application of quorum quenching (QQ) enzymes as a tool of probiotics. Only one example of digestive enzymes from the marine QQ probiotic candidate, Flaviramulus ichthyoenteri Th78T, will be presented in Table 1. QS is a population-dependent mechanism for bacteria to communicate, regulate gene expression, and synchronize social behaviors such as biofilm formation, bioluminescence, and secretion of virulence factors (Waters & Bassler, 2005). The enzymatic interruption of QS, termed QQ, has been suggested as a promising alternative antivirulence approach in plants and animals. The application of QQ bacteria in aquaculture as probiotics to control disease, which can be caused by Edwardsiella tarda, Aeromonas spp., and Vibrio spp., etc., has been proposed in several studies already. Several positive effects on larval survival of brine shrimp and turbot were demonstrated upon the addition of microbial enrichment cultures with QQ activities (Tinh, Yen, Dierckens, Sorgeloos, & Bossier, 2008; Van Cam et al., 2009). However, the majority of the reported QQ bacteria are of terrestrial origin, which would limit the application in aquaculture. Therefore, bacteria isolated from marine origins with QQ enzyme production activities will be mostly promising probiotic candidates in aquaculture. Most of the identified QQ enzymes are N-acyl-homoserine lactone (AHL)-degrading catalysts, which are responsible for the degradation of AHL as a population QS signal (Tang & Zhang, 2014). AHL-degrading

Table 1 Marine Microbial Enzymes as Tools in Probiotics to Benefit Animal Health Heat No. Enzyme Marine microbe Origin Resistant Substrate Specificity

References

Quorum quenching (QQ) enzymes 1

AHL lactonase FiaL

Flaviramulus ichthyoenteri Th78T

Flounder intestine

2

AHL lactonase MomL

Muricauda olearia Th120

Flounder Heat resistance skin mucus

AHLs (C6 to C14-HSL and 3-oxo-C6 to 3-oxo-C14-HSL)

Tang et al. (2015)

3

Unknown AHLdegrading QQ enzyme

Tenacibaculum discolor T84

Flounder Heat gill resistance

AHLs (C6 to C14-HSL and 3-oxo-C6 to 3-oxo-C14-HSL)

Tang et al. (2013)

4

AHL lactonase

Tenacibaculum soleae T133, T134, T148, T173, T189, T195

Flounder Heat gill resistance

AHLs (C6 to C14-HSL and 3-oxo-C6 to 3-oxo-C14-HSL)

Tang et al. (2013)

5

AHL lactonase

Olleya marilimosa T168 Flounder Thermolabile AHLs (C6 to C14-HSL gill and 3-oxo-C6 to 3-oxo-C14-HSL)

Tang et al. (2013)

6

Unknown AHLdegrading QQ enzyme

Colwellia aestuarii T171 Flounder Thermolabile AHLs (C8, C10, gill 3-oxo-C10, C12, 3-oxo-C12, C14, 3-oxo-C14)

Tang et al. (2013)

Zhang et al. (2016)

7

Unknown AHLdegrading QQ enzyme

Salinimonas lutimaris T194

Flounder Thermolabile AHLs (C8, C10, gill 3-oxo-C10, C12, 3-oxo-C12, C14, 3-oxo-C14)

Tang et al. (2013)

8

Unknown AHLdegrading QQ enzyme

Thalassomonas agariperforans-like T202

Flounder Thermolabile AHLs (C8, C10, gill 3-oxo-C10, C12, 3-oxo-C12, C14, 3-oxo-C14)

Tang et al. (2013)

9

Unknown AHLdegrading QQ enzyme

Rhodobacter ovatus J Th15

Flounder Thermolabile AHLs (C8, C10, C12, intestine C14, 3-oxo-C14)

Tang et al. (2013)

10 AHL lactonase

Flaviramulus basaltislike Th26, Th78, Th87, Th93, Th106

Flounder Heat intestine resistance

Tang et al. (2013)

11 Unknown AHLdegrading QQ enzyme

Marivita byusanensislike Th30

Flounder Thermolabile AHLs (C6, C12, C14) intestine

12 Unknown AHLdegrading QQ enzyme

Muricauda olearia Th120

13 Unknown AHLdegrading QQ enzyme

Pseudoalteromonas prydzensis Th125

AHLs (C6 to C14-HSL and 3-oxo-C6 to 3-oxo-C14-HSL)

Tang et al. (2013)

Tang et al. (2013)

Flounder Thermolabile AHLs (C10, 3-oxo-C10, Aranda et al. (2012) C12, 3-oxo-C12, C14, skin 3-oxo-C14) mucus Continued

Table 1 Marine Microbial Enzymes as Tools in Probiotics to Benefit Animal Health—cont’d Heat No. Enzyme Marine microbe Origin Resistant Substrate Specificity

References

14 AHL acylase

Tenacibaculum maritimum NCIMB2154(T)

AHLs (C10-HSL)

Romero, Avendan˜oHerrera, Magarin˜os, Ca´mara, and Otero (2010)

15 AHL lactonase AiiA

Bacillus cereus Y2

Marine source

AHLs

Lu, Yuan, Xue, Zhang, and Zhou (2006)

16 Alginate lyases and lipases

Flaviramulus ichthyoenteri Th78T

Flounder intestine

Zhang et al. (2016)

17 Enzymes for production of B vitamins

Flaviramulus ichthyoenteri Th78T

Flounder intestine

Zhang et al. (2016)

Flaviramulus 18 Sialic acid lyases, sialidases, sulfatases, ichthyoenteri Th78T and fucosidases

Flounder intestine

Zhang et al. (2016)

Digestive enzymesa

a

Other digestive enzymes from marine probiotics have been reviewed by Nguyen et al. (2013) and Nguyen (2016).

Characterization and Applications of Marine Microbial Enzymes

61

enzymes may be classified into three major types according to their enzymatic mechanisms: AHL lactonase (lactone hydrolysis), AHL acylase (amidohydrolysis), and AHL oxidase and reductase (oxidoreduction) (Tang & Zhang, 2014). AHL lactonase hydrolyzes the ester bond of AHL yielding the corresponding N-acyl-homoserine. This hydrolyzation may also occur spontaneously at alkaline pH and may be reversed under acid pH. AHL acylase hydrolyzes the amide bond of AHL to yield a homoserine lactone and the corresponding fatty acid chain, whereas AHL oxidase and reductase usually catalyze a modification of AHLs. In most cases, AHL lactonases require metal ions (except AiiM and QsdH) and target both shortand long-acyl chain AHLs. Unlike lactonases, acylases exhibit substrate specificity based on the length of the acyl chain and the substitution on the β-position of the AHL chain. The first report of the degradation of AHL acylase activity in a QS bacterium and HSL lactonase activity in any bacterium came from a soil pseudomonad and P. aeruginosa PAO1 (Huang, Han, Zhang, & Leadbetter, 2003). Enzymatic degradation of AHLs has been extensively studied and found in many organisms including mammals, plants, fungi, archaea, and bacteria (Draganov et al., 2005; Lasarre & Federle, 2013; Tang et al., 2013), although the genes responsible for AHL-degrading activity in plants and fungi have not been identified. One of the first examples of marine QQ strains was Bacillus cereus Y2 with the capability of inactivating lactonase-type AHLs (Lu et al., 2006). In other way, one of the first acylase-type marine enzymes came from Tenacibaculum maritimum, a filamentous, biofilm-forming member of the Bacteroidetes, which causes the widely distributed marine fish disease tenacibaculosis. In the strain NCIMB2154(T), a degradation activity for long-acyl AHLs (C10-HSL) was demonstrated (Romero et al., 2010). Currently, diverse marine QQ strains have been identified from microbiota of gill, intestine, and skin mucus of healthy flounder. The epidermal mucus plays an important role in the fish innate immune system (Subramanian, MacKinnon, & Ross, 2007), and the adhesion to mucus surfaces has been considered as a necessary criterion for probiotics (Chabrillon, Rico, Balebona, & Morinigo, 2005). For example, a mucus-derived strain, Pseudoalteromonas prydzensis Th125, has recently been demonstrated to express antibacterial activity against several vibrios (Aranda et al., 2012). Based on the properties of P. prydzensis Th125, it may become potential probiotics to control disease in aquaculture. However, QQ enzymes have been not uncovered, obtained, or purified to explore probiotic properties and mode of actions in this strain.

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Using a simple, sensitive, and high-throughput method based on the biosensor Agrobacterium tumefaciens A136, other 25 QQ bacterial strains belonging to 14 species were identified from 366 candidate bacterial strains from healthy flounder (Tang et al., 2013). Twenty of 25 bacterial strains were typical marine bacteria, except 5 strains with high similarity to Sphingopyxis spp. and Novosphingobium tardaugens. Twelve of the 14 bacterial species have not been reported to bear AHL-degrading activities yet in the previous studies. Similarly, 24 QQ strains of 106 isolates from various marine environments were identified with weak AHL-degrading activities (Romero, MartinCuadrado, Roca-Rivada, Cabello, & Otero, 2011). Especially, the lactonases and the heat-resistance of QQ activities, which were proposed when C6-HSL was used as the substrate, were demonstrated in three species Tenacibaculum soleae, Olleya marilimosa, and Flaviramulus basaltis-like species (96.7–96.8% identities in 16S rRNA gene sequences to F. basaltis H35T) (Table 1) (Tang et al., 2013). The results indicated that different types of QQ enzymes present in different species of Tenacibaculum or even in different strains of Tenacibaculum discolor. Also, it was possible that there were different enzymes (e.g., lactonase and acylase) existing in different strains of O. marilimosa (Tang et al., 2013). Among them, strain Th78T was identified as F. ichthyoenteri (Zhang et al., 2016) and demonstrated to degrade lactonase-type AHLs. The degrading activity of lactonase in this strain exhibited substrate specificity with all tested AHLs of different acyl chains including C6-homoserine lactone (HSL), 3-oxo-C6-HSL, C8-HSL, 3-oxo-C8-HSL, C10-HSL, 3-oxo-C10-HSL, C12-HSL, 3-oxo-C12-HSL, C14-HSL, and 3-oxo-C14-HSL (Tang et al., 2013). The name FiaL (F. ichthyoenteri N-acyl-homoserine lactonase) was proposed as a novel marine AHL lactonase encoded by gene GL001211 based on relatively low identity with the reported AHL lactonases (Zhang et al., 2015). Interestingly, FiaL homologs can be found in various marine bacteria of the family Flavobacteriaceae. FiaL shows the highest identity (70%) to a predicted protein in Tenacibaculum ovolyticum (WP_028890039), followed by Polaribacter sp. Hel_I_88 (WP_026776245, 63% identity) and Aquimarina muelleri (WP_027414219, 61% identity). However, the QQ activity of these homologous proteins remains to be confirmed experimentally. AHL lactonases and acylases are two most popular types of QQ enzymes, and lactonases normally presented broad AHL-inactivating activity (Dong et al., 2001), while many acylases were specific to long-chain AHLs (Lin et al., 2003). Previous observations indicated that long-chain AHLdegrading activities might be more popular among marine cultivable

Characterization and Applications of Marine Microbial Enzymes

63

bacteria (Romero et al., 2011; Tang et al., 2013) and also among marine metagenome samples (Romero, Martin-Cuadrado, & Otero, 2012); thus, AHL acylases might be more common than lactonase in the ocean. Besides the degradation of AHL as the most common QS signal, QQ enzymes are also involved in the degradation of other interspecies signaling molecules such as diffusible signal factor (DSF), 2-heptyl-3-hydroxy-4(1H)quinolone (PQS), and autoinducer-2 (AI-2) (Tang & Zhang, 2014). DSF is a fatty acid signal molecule involved in the regulation of virulence. Several bacterial strains belonging to genera Bacillus, Paenibacillus, Microbacterium, Staphylococcus, and Pseudomonas were identified that were capable of particularly rapid degradation of DSF. However, the rapid degradation of DSF has not been detected in these bacteria. The typical example, carAB, which was required for the synthesis of carbamoyl phosphate, a precursor for pyrimidine and arginine biosynthesis, was required for the rapid degradation of DSF in Pseudomonas spp. strain G (Newman, Chatterjee, Ho, & Lindow, 2008). In addition, the 2,4-dioxygenase, Hod, involved in the quinaldine utilization pathway in Arthrobacter nitroguajacolicus is able to cleave PQS. In P. aeruginosa PAO1 cultures, exogenously supplied Hod protein reduced the expression of the PQS biosynthetic gene pqsA, expression of the PQS-regulated virulence determinants lectin A, pyocyanin, and rhamnolipids, and virulence in plants. This indicates that enzyme-mediated PQS inactivation has the potential as an antivirulence strategy against P. aeruginosa (Pustelny et al., 2009). Finally, when LsrK is artificially provided in vitro, the extracellular phosphor-AI-2 molecules cannot be transported into cells and are degraded overnight. LsrK-mediated degradation of AI-2 attenuates the QS response in S. enterica serovar Typhimurium and Vibrio harveyi even though the AI-2 signal transduction mechanisms and the phenotypic responses are species specific (Roy, Fernandes, Tsao, & Bentley, 2010). Therefore, the enzymatic approach for quenching QS systems will spawn new methods for controlling cell phenotype and potentially open new avenues for controlling bacterial pathogenicity. Unfortunately, non-AHL degradative enzymes have been not reported from QQ bacteria of marine origin.

5. CONCLUSIONS Marine microorganisms provided an almost untapped reservoir of diverse novel and useful enzymes. Because of a restriction in exploration, a large proportion of these organisms have not been identified and investigated for enzyme activity. This review presents the purification,

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characterization, and applications of marine microbial enzymes in biotechnology and probiotic technology. Microbial enzymes of marine origin can be used a tool for biotechnological processes in a series of industrial fields such as dairy, food, detergents, textile, pharmaceutical, cosmetic, and biodiesel production, and novel chemical and material synthesis. Additionally, this review describes the potential applications of QQ enzymes from probiotics to control pathogens in marine aquaculture. Hopefully, these novel approaches can serve as a reference for assessing enzymes from marine environments as a valuable tool in technology development and innovation.

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CHAPTER FOUR

Biotechnological Applications of Marine Enzymes From Algae, Bacteria, Fungi, and Sponges S. Parte1, V.L. Sirisha, J.S. D’Souza2 UM-DAE Centre for Excellence in Basic Sciences, Mumbai, India 2 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 1.1 Oceanic Habitat and Marine Ecosystem 1.2 Marine Biotechnology 2. Enzyme Technology: Current State of the Art 3. Marine Enzymes 4. Biotechnological Applications and Advantages of Enzymes From Marine Algae, Bacteria, Fungi and Sponges 4.1 Algae 4.2 Bacteria 4.3 Fungi 4.4 Sponges 5. Challenges Encountered in Harnessing Marine Resources 6. Future Prospects References

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Abstract Diversity is the hallmark of all life forms that inhabit the soil, air, water, and land. All these habitats pose their unique inherent challenges so as to breed the “fittest” creatures. Similarly, the biodiversity from the marine ecosystem has evolved unique properties due to challenging environment. These challenges include permafrost regions to hydrothermal vents, oceanic trenches to abyssal plains, fluctuating saline conditions, pH, temperature, light, atmospheric pressure, and the availability of nutrients. Oceans occupy 75% of the earth’s surface and harbor most ancient and diverse forms of organisms (algae, bacteria, fungi, sponges, etc.), serving as an excellent source of natural bioactive molecules, novel therapeutic compounds, and enzymes. In this chapter, we introduce enzyme technology, its current state of the art, unique enzyme properties, and the biocatalytic potential of marine algal, bacterial, fungal, and sponge enzymes that have 1

Current address: Postdoctoral Research Associate, Department of Physiology, CTR Bldg, Health Sciences Campus, University of Louisville, Louisville, KY 40202, United States.

Advances in Food and Nutrition Research, Volume 80 ISSN 1043-4526 http://dx.doi.org/10.1016/bs.afnr.2016.10.005

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indeed boosted the Marine Biotechnology Industry. Researchers began exploring marine enzymes, and today they are preferred over the chemical catalysts for biotechnological applications and functions, encompassing various sectors, namely, domestic, industrial, commercial, and healthcare. Next, we summarize the plausible pros and cons: the challenges encountered in the process of discovery of the potent compounds and bioactive metabolites such as biocatalysts/enzymes of biomedical, therapeutic, biotechnological, and industrial significance. The field of Marine Enzyme Technology has recently assumed importance, and if it receives further boost, it could successfully substitute other chemical sources of enzymes useful for industrial and commercial purposes and may prove as a beneficial and ecofriendly option. With appropriate directions and encouragement, marine enzyme technology can sustain the rising demand for enzyme production while maintaining the ecological balance, provided any undesired exploitation of the marine ecosystem is avoided.

1. INTRODUCTION With the ever-increasing human population, the demand for basic needs (food, shelter, clothing, medicine, etc.) has risen immensely. Simultaneously, a phenomenal upsurge of technological advancement and industrialization has inflicted tremendous pressure on natural resources. Marine habitat offers diverse ecosystems and serves as an excellent source of natural bioactive molecules, novel compounds, secondary metabolites, enzymes, etc. Investigating their varied applications to develop several biotechnological products and services is an attractive strategy. This chapter summarizes current state of the art in enzyme technology for those available from marine organisms and updated information about their diverse biotechnological applications and functions, encompassing sectors such as domestic, industrial, commercial, and healthcare.

1.1 Oceanic Habitat and Marine Ecosystem Oceans occupy 75% of the earth’s surface and harbor most ancient and diversified life forms (Thakur & Thakur, 2006). Ocean floors constitute unique features such as permafrost regions, hydrothermal vents, cold seeps, oceanic trenches, abyssal plains, extreme saline conditions, fluctuating temperatures (4–100°C or 1.5 to 350°C), pressures (1–100 atm.), variable/ deficient light (highly photic to nonphotic zones), and nutrient conditions (oligotrophic to eutrophic). Marine habitat upholds titanic treasures of resources in terms of diverse flora–fauna (300,000 species) of which only a small proportion have been explored (Malakoff, 1997; Winston, 1988).

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The quest for exploring marine/oceanic habitat for resources began in the 1960s as researchers realized the true potential of the abundant treasures offered. Of the 34 known animal phyla, 19 are present in the oceans, the other 9 occupy both land and sea, while 2 are terrestrial (Raghukumar, Damare, & Singh, 2010). Out of 300,000 species, >10,000 marine metabolites have been explored, isolated, and characterized (Bhatnagar & Kim, 2010; Davidson, 1995). Since marine organisms have evolved to thrive in diverse and extreme challenging conditions, they exhibit a remarkable level of specialization, and their secondary metabolites especially secreted enzymes may serve as better prospects for industrial and biomedical purposes. Marine organisms produce metabolites having a deep influence as antitumor, antiviral, and enzyme inhibitory agents; these affect central nervous, respiratory, neuromuscular, autonomic nervous, cardiovascular, and gastrointestinal systems, while other bioactive properties are related to drug discovery (Bonugli-Santos et al., 2015; Imhoff, Labes, & Wiese, 2011; Nikapitiya, 2012). Bioactive secondary metabolites, anticancer agents, etc., from marine habitat have been explored from diverse classes of organisms and contribute in variable proportion such as algae–microalgae (cyanobacteria and diatoms, 9%), bacteria (18%), and sponges (37%). A general shift in preference toward using natural products vs synthetically manufactured ones is observed, and marine resources are being explored in food, pharmaceuticals, healthcare, biomedicine, nutraceuticals and cosmeceuticals. Following sections will unfold how the process of harnessing marine biodiversity did not remain untouched by the recent biotechnology boom and brought about a revolution in utilizing marine ecosystem for the development of enzyme technology in particular.

1.2 Marine Biotechnology Marine biotechnology is a multidisciplinary science involving the application of marine organisms and engineering them for products or services that benefit mankind. It was recently recognized that the level of interdisciplinary research was insufficient and immense networking was the need of the hour, giving rise to “The Marine Biotechnology ERA-NET (ERA-MBT)” consortium of funding agencies from 19 European nations in December 2013, whereby they pool and share joint funds, resources, and undertake several developmental projects in Marine Biotechnology on a nationwide scale based on core objective of Europe 2020 strategy of the EU. The longterm goal is the basic research using expertise of marine biologists, marine

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invertebrate zoologists, taxonomists, plant physiologists, phycologists, microbiologists, genome scientists, chemists, bioengineers (bioprocess engineers), chemical engineers, biotechnologists, nanotechnologists, systems biologists, bioinformaticians, toxicologists, -omics technologists, etc. Biotechnological applications in terms of developing novel drugs, hormones, vaccines, crops, etc., broadly fall under areas of biomedical, agricultural, environmental, and industrial biotechnology. Biotechnology boom in several fields such as medicine, agriculture, and food has already occurred, and only recently the marine biotechnology revolution has made a huge impact. Marine biotechnology finds its applications in multiple areas such as human and environmental health, bioremediation, biomining, biofouling, biomass, food production, biomedical products, organic compounds, aquaculture, cell culture, fishery, bioprospecting, marine molecular, biomaterial and bioprocess biotechnology, genomics, bioinformatics, and nanotechnology while utilizing marine biological resources. Other fields include production of fine chemicals and agrichemicals, food supply, cosmetics, pharmaceuticals, nutritional supplements/nutraceuticals, and molecular probes for the production of commercially important enzymes (Pomponi, 1999; Ritchie, Guy, & Philp, 2013; Thakur & Thakur, 2006; Tramper et al., 2003). Species diversity and unique features of marine ecosystem (such as deep oceanic regions, hydrocarbon seeps, hydrothermal vent regions, freezing temperature zones, regions away from sea coast within low-nutrient region, or with higher nutrient levels near seashores with symbiotic and commensal-type association between flora and fauna, high pressure, and salinity regions) enhance novel bioactive chemical compound production and biotechnologically relevant processes such as anaerobic biotransformation, bioremediation, biodegradation, and biohydrometallurgy. Such conditions foster production of antifreeze compounds, surfactants, cold-active enzymes, and novel bioactive chemical compound production related to sensing and signaling defense to name a few which could be further explored (Glover et al., 2010; Rogers et al., 2012). Marine biotechnology may thus contribute immensely toward the world of biotechnology market in future if the resources are explored to their maximum potential. Certain challenges such as development of identification/ screening technology, isolation, optimization of production, recovery, and maintenance of sustained supply of marine bioproducts for various purposes persist in this field and will be discussed further. A sound understanding of various cellular and molecular processes regulating production of these commercially important metabolites is lacking and needs to be explored in depth.

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Precise knowledge about the mechanism of action of microbial metabolites especially enzymes with a close association between marine biology and microbial biotechnology while harnessing the understanding of various biogeochemical cycles and their synchronized interactions for developing marine enzyme technology is the need of the hour.

2. ENZYME TECHNOLOGY: CURRENT STATE OF THE ART Enzymes are employed at industrial scale in manufacturing of food (bread, cheese, beer, and vinegar), fine chemicals (amino acids, vitamins), agriculture (growth hormones, cytokinins), and pharmaceuticals (insulin). Although enzymes can be obtained from several organisms such as microorganisms, plants, and animals, microorganisms prove to be the best and most reliable source. This is attributed to their biochemical diversity, ease of culturing for large-scale production, their commercial applications, genetic manipulation, and superior quality with remarkable stability and activity as compared to their terrestrial counterparts. Of the known 4000 enzymes, only 5% are of bacterial origin and are used commercially; only 10% of these are used at industrial scale. Although 12 major and 400 minor producers/suppliers of enzymes are known worldwide, almost 75% are produced by enzyme companies, such as Novozymes (Denmark), DuPont (USbased currently acquired by Denmark-based Danisco), and Roche (Switzerland). Majority of enzymes are utilized in food industry, where 75% of all industrially used enzymes are hydrolytic in nature such as amylase, protease, lipase, cellulase, xylanase, catalase, and laccase/ligninase dominating the enzyme market (Li, Yang, Yang, Zhu, & Wang, 2012). Conventional enzymes catalyze specific reactions; however, improving their quality and efficiency for catering to diverse applications is the main concern. In order to boost the bioprocessing technology industry, specially designed or engineered enzymes have become biocatalysts of choice over chemically modified ones, thereby adding to the versatility and usefulness of products in global market. In order to develop cost-efficient production of microbial enzymes at industrial scale, several key factors (selection of host organism, substrate involved in enzyme production, environmental conditions, etc.) need to be considered, while mere knowledge about their typical chemical and stereochemical properties which impart them a cutting edge over the terrestrially available ones and interaction between various factors is not sufficient. Characteristic properties of marine enzymes befit their use for biotechnological and industrial applications and attract further research

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(Trincone, 2011). Recent advances in recombinant gene technology, metagenomics, whole genome shot gun sequencing, directed evolution, bioinformatics, transcriptomics, active-site imprinting, de novo protein designing, proteomics, developing novel platforms for discovering new protein catalysts used in industry, developing novel substrates for known enzymes, enzyme immobilization, etc., have aided in the development of improved and cheaper biocatalytic enzymes, highly efficient for wider scope of applications. These rationally designed enzymes exhibit advanced properties compatible with dynamic or robust scale biocatalytic processes such as improved activity, thermal and solvent stability, substrate and/or product specificity and tolerance, and enantio-, chemo-, regio-, and stereoselectivity. Immobilization of enzymes or coimmobilization of multienzyme systems (activated by different reactive groups), enzyme-aided immobilization of enzyme, and development of fusion proteins or their photoimmobilization (by exposure to UV/sunlight) onto respective support surfaces or carrier materials take into account several parameters. These include protein/support attachment, mass transference (enhanced by microwave irradiation), modulation of immobilization or distribution rate, etc., which determine the success of the procedure (Li et al., 2012; Venter, 2004; Verma, Chaudhary, Tsuzuki, Barrow, & Puri, 2013).

3. MARINE ENZYMES The serious drawbacks of chemical catalysis are cost-ineffectiveness due to nonspecific reactions resulting into undesirable by-products, requirements of high temperature, pressure and resultant large volumes of cooling water downstream, dedicated and sophisticated equipment, and detrimental effects on human health and environment. An immediate need for utilizing biocatalytic agents possessing less toxic/hazardous effects was felt. This marked the beginning of the era of Marine Microbial Biotechnology and subsequent positive repercussions in its stride. An exclusive field known as Marine Enzyme Technology has come into existence where enzymes obtained from marine organisms serve as best contenders due to their advantageous characteristics and diverse applications (van Beilen & Li, 2002). Use of enzymes demanded relatively simpler requirements to set up the reactions and in turn provided highly specific and high reaction rates in terms of productivity. It meant utilization of algae, bacteria, fungi and sponges of marine origin for the manufacture of commercial products (described in Tables 1–4).

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Marine enzymes are isolated from organisms dwelling in saline/brackish environment and possessing diverse applications and functions to accelerate bioprocesses efficiently. Fermentation process employed natural renewable resources while applying least pressure against and thus favorably sustaining the vulnerable environment (Sarkar, Pramanik, Mitra, & Mukherjee, 2010). Enzyme-driven mild operating conditions have enabled easy control over the processes and simplified production steps. Relatively lesser amounts of enzyme preparations (solid or liquid) could be employed in the system, thus reducing their cost, storage space, and release of waste effluents into the environment. Bioprocess engineering for marine enzyme production involves development of batch, fed-batch, continuous, and shake flask cultures. Submerged, immobilized, and solid-state processes are included in batch cultures, whereas continuous culture comprises suspended and immobilized cultures (Sarkar et al., 2010). Improvised technology such as immobilized enzymes enables cost-cutting involved in production step as they allow reusage, are stable, reduce reaction time, improve better control over the process, easy to separate, and hence less labor involved (KatchalskiKatzir & Kraemer, 2000). Genetically modified microorganisms can specifically improve quality of enzyme preparation such as enhance tolerance at high temperature, stability in extreme environmental conditions such as fluctuating pH (acidic or alkaline) conditions, maintaining optimum activity during high substrate concentrations to achieve maximum productivity, or the presence of agents like metal ions and other compounds for mass production of commercially important enzymes. Resistant microbial strains, e.g., thermostable or thermophilic organisms, offer several advantages over the conventional host strains utilized for enzyme production such as contamination from other nonthermotolerant bacteria is minimized at higher temperatures, enabling reactions to last for longer duration without interference from nonspecific strains of organisms. It also permits proper breakdown of substrate and release of raw materials used, and aids in proper penetration of enzyme by enhanced mass transfer and reduction in its viscosity, thus improving the rate of reaction and product formation, e.g., xylanase-a hydrolytic enzyme used in brewing at an industrial scale, laccase, and its xerophytic isoforms useful in processes like textiles, dyeing, pulping, and bioremediation. Site-directed mutagenesis and directed evolution have been employed for enhancing thermal stability of known enzymes, while a combination of both strategies is popularly being adopted among researchers. Molecular modeling may be employed for rational prediction of protein secondary and tertiary structures based on known amino acid sequence. Thus,

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major enzymes with commercial significance (cellulases, amylases, lipases, xylanase, proteases) serve as an ecofriendly and profitable alternative to chemical catalysis for large-scale utility (Nigam, 2013). Marine microbial biodiversity has aided the development of biomedical fields. Harsh marine environment, extreme conditions, and strong selection due to process of evolution have been known to impart a characteristic diversity and extraordinary biocatalytic potential to marine enzymes obtained from micro- and macroorganisms and are described further. This has provided great impetus to marine microbial enzyme technology to prosper and revolutionize the bioprocesses which were earlier explored by redundant methods. Moreover, application of genetic and protein engineering has improved efficiency and productivity of enzymes and is now being employed for novel processes, and hence several countries like Denmark, Switzerland, Germany, Netherlands, and United States have emerged as major contributors in the global enzyme market (Li et al., 2012).

4. BIOTECHNOLOGICAL APPLICATIONS AND ADVANTAGES OF ENZYMES FROM MARINE ALGAE, BACTERIA, FUNGI AND SPONGES As mentioned earlier, enzymes produced by marine organisms (algae, bacteria, fungi and sponges) exhibit unique physiological properties such as hyperthermostability, barophilicity, salt and pH tolerance, adaptivity to extreme cold conditions, and novel chemical and stereochemical properties. Unlike their terrestrial counterparts, these properties enable them to catalyze chemical reactions under extreme conditions (Saleem et al., 2007; Zhang & Kim, 2010) and confer upon them unique potential for tremendous biotechnological applications (Tables 1–4). In order to utilize them for diverse array of industrial and biotechnological applications, these enzymes ought to be in the purest form possible. For this purpose, several techniques in a combinatorial and sequential manner have been employed till date to recover enzymes from marine sources.

4.1 Algae Marine algae also known as seaweeds are macroscopic and multicellular in nature, and lie at the base of the food web and include members of the red (Rhodophyta), brown (Phaeophyta), and green (Chlorophyta) algae, providing shelter for numerous fishes, invertebrates and mammals. Seaweeds/marine algae are a food source for humans; food additives such

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as alginates and carrageenan are used in cooking, baking, and medicine. Alginates are used in wound dressings, for production of dental molds and agar (bacterial growth media). Seaweeds are ingredients in toothpaste, cosmetics, and paints, and are used to manufacture products such as adhesives, dyes, gels, paper coatings, and explosives. Algal metabolites such as carbohydrates, proteins, vitamins, carotenoids, and lipids are used in medicine, food and feed additives, cosmetics, and for energy production. Microalgae are known as living cell factories as they can produce biofuels and various other bioactive compounds useful in food, aquaculture, poultry and pharmaceutical industries. Green algae (Chlorophyceae), Chlorella vulgaris, Haematococcus pluvialis, Dunaliella salina, Cyanobacteria, and Spirulina maxima are the most biotechnologically significant species of microalgae and are successfully employed in the manufacture of nutritional supplement and animal feed additives. Brown, red, and green algae also provide resources like pigments, minerals, polyphenols, and trace elements oils. Several enzymes with medicinal, pharmaceutical, and biotechnological relevance (described in Table 1) are produced by marine algae such as Mannuronan C5 epimerase with functions similar to alginate lyase by the brown algae Saccharina japonica (Inoue et al., 2016); α-1,3-glucosidase is produced by red microalga Porphyridium sp. (Rhodophyta) (Levy-Ontman et al., 2015); some extra- and intracellular enzymes produced by Cylindrotheca, Closterium, D. salina, and Chaetoceros muelleri species (Gao & Chi, 2015) are involved in the biodegradation of diethyl phthalate (which causes toxicity and affects androgen biosynthesis). The enzymes SOD, ascorbate peroxidase, glutathione peroxidase and catalase are directly involved in conferring metal tolerance and hence implicated in bioaccumulation and as useful biosensors for marine pollution (Babu et al., 2014; Moenne et al., 2016). Symbiotic association between seaweeds and bacteria are a classic example during manufacturing of alginate from brown algae where some depolymerization occurs due to physical forces as well as by alginate lyases produced by bacteria associated with the seaweeds (Ertesva˜g, 2015).

4.2 Bacteria Marine bacteria are single-celled organisms constituting the marine microbiome accounting >98% of the ocean biomass. Adaptation to the diverse marine environment is an exclusive feature of marine organisms and enzymes confer exceptional abilities to thrive successfully in such adverse environments (Zhang & Kim, 2012). As a consequence, multifunctional

Table 1 Details of Marine Enzymes Derived From Algae Enzyme Enzyme Source

Enzyme Function and/or Applications

Reference

Mannuronan C5 epimerase with Brown algae Saccharina japonica Required for polymerization, epimerization, and functions similar to alginate lyase biosynthesis of alginate polysaccharide implicated for industrial usage

Inoue et al. (2016)

α-1,3-Glucosidase, glucosidase II Red marine microalga (GANAB) glycoenzyme (NAD+- Porphyridium sp. (Rhodophyta) dependent D-lactate dehydrogenase; glycolate oxidase)

Levy-Ontman, Fisher, Shotland, Tekoah, and Arad (2015)

Required for polysaccharide biosynthesis which has applications as stabilizers in food industry; for production of other industrial, medicinal, and pharmaceutical compounds

Tocher and Meeuse Implicated in transport of water and help in biological p-Coumaryl alcohol biosynthesis Green and red algae, Monostroma fuscum (Postels and defense for the uni- and multicellular algae; derivatives of (1966) and Labeeuw, enzyme, phenolaseMartone, Boucher, p-coumaryl alcohol act as antioxidants in food industry Ruprecht) oxidoreductase and Case (2015) Extra- and intracellular enzymes Cylindrotheca, Closterium, Dunaliella salina, Chaetoceros muelleri

Biodegradation of diethyl phthalate (DEP) and di-n-butyl Gao and Chi (2015) phthalate (DBP) as phthalates cause chronic toxicity and inhibit androgen biosynthesis

Squalene synthase-like enzyme

Botryococcus braunii Race L

Squalene synthase (SS) forms C30 squalene, a precursor for Thapa et al. (2016) sterols/steroids, bile acids, lipoproteins, and vitamin D biosynthesis in eukaryotes and hopanoids in some prokaryotes; geranylgeranyl pyrophosphate (GGPP), an intermediate in the process is implicated in lytic bone disease and other disorders—nonskeletal cancers, neurodegenerative, and cardiovascular diseases

SOD, ascorbate peroxidase, glutathione peroxidase, catalase

Marine red and green macroalga—Rhodophyta and Chlorophyta; Acanthophora spicifera, Chaetomorpha antennina, Ulva reticulata

Responsible for activation of antioxidant system, confers Babu et al. (2014) and Moenne, Gonza´lez, extreme tolerance to copper metal useful in metal and Sa´ez (2016) bioaccumulation, useful as bioindicators of marine pollution

Biotechnological Applications of Marine Enzymes

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enzymes have been explored, and recently, the enzyme Amy63 from South China sea was reported to be isolated from Vibrio alginolyticus possessing amylase, agarase, and carrageenase activities (Liu, Wu, Jin, & Sun, 2016). They also perform multiple functions, which differ mechanistically and are implicated in biomedical and fundamental research related to diseases (Huberts & van der Klei, 2010). Proteases constitute 60% of all industrial enzymes required in diverse applications such as detergent, leather, and in pharmaceutical industry for digestive and antiinflammatory drugs. Among the hydrolases, lipases are important in the detergent industry and occupy 80–90% of the enzyme detergent market. They are also required for production of paper, cosmetics, food flavoring, organic synthesis, and other industrial applications (Zhang & Kim, 2012). Esterase can be used for hydrolytic cleavage of polyethylene and degradation of plastic, synthesis of esters, preparation of optically active compounds, food, textile industries, and deinking (Kulkarni, Patil, & Satpute, 2013). Deinking process may be catalyzed by enzymes like cellulases, xylanases, esterases, lipases, and lignolytic enzymes, and marine enzymes need to be explored. Marine bacterial enzymes include chitinase, chitosanase, alginate lyase, agarase, carrageenase, cellulose and hemicellulose hydrolase, xylanase, DNA polymerase, ligase, and restriction endonuclease from extremophilic bacteria (psychrophilic, thermophilic, acidophilic, alkaliphilic, halophilic; Zhang & Kim, 2012). Of specific importance are the cold-adapted and surfactant-stable alginate lyase, thermostable DNA polymerase from Thermus aquaticus (Taq), thermostable alkaline protease, halotolerant thermostable lipase, novel halo-alkali-tolerant and thermotolerant chitinase, cold-active and salt-tolerant α-amylase (AmyZ), xylanase, cold-active endo-β-1,4-xylanase, cold-adapted phosphinothricin N-acetyltransferase enzymes (Table 2) which function in nonconventional conditions with optimal specificity, thus curbing high costs involved in industrial manufacturing processes to maintain temperatures, pH, pressure, etc.

4.3 Fungi Fungi are eukaryotic heterotrophs existing as single filaments or aggregates in almost all sorts of niches such as oceans, coastal areas, estuaries, on land, or mangrove swamps. Results of high-throughput sequencing methods predict 5.1 million fungal species to exist on earth, of which marine fungi comprise >1500 species. Marine fungi thrive in the sea either as obligates or in facultative modes, exist as free-floating entities or lodge onto driftwood, sand

Table 2 Details of Marine Enzymes Derived From Bacteria Enzyme Enzyme Source

Enzyme Function and/or Applications Reference

A-amylase has implications in food, Huberts and van der Klei (2010) and Liu et al. (2016) pharmaceutical, and chemical industry; multifunctional amylase exhibits transglycosylation and hydrolysis activities to produce isomaltooligosaccharides and maltooligosaccharides and glucose, multifunctional enzymes are implicated in diseases

Amy63 with amylase, agarase, and carrageenase activities

Vibrio alginolyticus

Proteases, amylases, cellulases, chitinases, keratinase, xylanases, nitrogen-assimilating enzyme, carboxymethylcellulase

Hemolytic and anticancer activity Citrobacter, Proteus, Bacillus, Corynebacterium sp., marine Actinobacteria, Bacillus safensis, Argyrolobium roseum, Bacteria from marine sedentary organisms, Streptomyces bikiniensis— ESS_amA-1 strain

Erythrobacter Glycosyl hydrolase, cytochrome P450 (CYP) and uridine diphosphoglucuronosyl, transferase, epoxide hydrolase (epoxide hydratase)

Useful for novel industrial applications

Vijayasurya, Gayathri Devi, Tintu, and Manjusha (2014), Manivasagan, Venkatesan, Sivakumar, and Kim (2013), Lateef, Adelere, and GueguimKana (2015), Habib, Zia, Bibi, Abbasi, and Chaudhary (2015), and Ahmad, Yasser, Sholkamy, Ali, and Mehanni (2015) Yaakop et al. (2015)

Cold-adapted and surfactant-stable alginate lyase; alginate acetylase, alginate lyase, alginate deacetylase

Sphingomonas sp. strain, A1 Cause degradation of polysaccharides to yield di- and Agarivorans sp. L11S; Pseudomonas, Azotobacter trisaccharides as the main products which have potential applications in industry for alginate biosynthesis

Takase, Mikami, Kawai, Murata, and Hashimoto (2014), Li, Yang, Zhang, Yu, and Han (2015), and Ertesva˜g (2015)

Thermophilic eubacterium Vent DNA polymerase, thermostable DNA polymerase from from deep-sea hydrothermal vent; isolated Thermus aquaticus (Taq) NS-E is an obligately anaerobic, extremely thermophilic, heterotrophic eubacterium

Enzyme used for polymerase chain reaction, DNA manipulation in vitro, useful for cloning, sequencing, labeling, mutagenesis, etc., implicating its significance in molecular biology/biotechnology studies

Belkin, Wirsen, and Jannasch (1986), Innis, Myambo, Gelfand, and Brow (1988), and Ishino and Ishino (2014)

Phosphoenolpyruvate carboxykinase Vibrio costicola (PEPCK), polyketide synthase

Biosynthesis of secondary metabolites—erythromycin, rapamycin, tetracycline, lovastatin, resveretrol

Salvarrey, Cannata, and Cazzulo (1989) and Thakur and Thakur (2006)

Thermostable alkaline protease

SD11 halophilic marine bacterium

Industrial applications

Cui, Wang, and Yu (2015)

Halotolerant thermostable lipase

Oceanobacillus sp. PUMB02 Disruption of bacterial biofilms

Seghal, Nishanth, Kennedy, Dobson, and Selvin (2014)

M12 metalloprotease myroilysin protease

Deep-sea bacterium Myroides profundi D25

Shao et al. (2015)

Elastinolytic activity, strong collagen-swelling ability, collagenolytic activity, good collagen modifier

Continued

Table 2 Details of Marine Enzymes Derived From Bacteria—cont’d Enzyme Enzyme Source

Enzyme Function and/or Applications Reference

Fucoidanase (FNase S)

Marine bacterium Sphingomonas paucimobilis PF1

Cause hydrolysis of fucoidan, thus Kim, Park, Park, Choi, and acting as a valuable tool for structural Park (2015) analysis of fucoidans, and production of bioactive fuco-oligosaccharides

Alkaline phosphatase, acid phosphatase, esterase (C4 and C8), and naphthol-ASBI-phosphohydrolase, leucine, valine, and cystine arylamidase, trypsin, α-chymotrypsin, acid phosphatase, α- and β-galactosidase, glucuronidase, α-glucosidase, β-glucosidase, N-acetyl-βglucosaminidase, α-mannosidase, α-fucosidase

Baltic cyanobacteria— Nodularia spumigena CCNP1401; marine bacterium Cobetia marina (strain KMM MC-296) isolated from coelomic liquid of the mussel Crenomytilus grayanus

May be useful to study structure and function of nucleic acids, for preparation of Ig-enzyme conjugates for immunologic assays; biofuel, pharmaceutical, production of fine chemicals, and food industries

Novel halo-alkali-tolerant and thermotolerant chitinase

Pseudoalteromonas sp. DC14 Halo-alkali and thermostable from Caspian sea properties are useful for diverse industrial applications, eg: useful for hydrolysis of chitin and hence N-acetyl chitobiose production which in turn is useful in fermentation research, biochemical enzyme assays, and in vitro diagnostic analysis

Mazur-Marzec et al. (2015), Dalmaso, Ferreira, and Vermelho (2015), and Yu et al. (2005)

Makhdoumi, DehghaniJoybari, Mashreghi, Jamialahmadi, and Asoodeh (2015)

Vibrio β-glucosidase, laminarinase

Vibrio campbellii

Endohydrolysis of 1,3- or 1,4linkages in β-D-glucans with implications in industrial usage

Wang et al. (2015)

Alkaline protease/alkaline serine protease

Bacillus circulans BM15, Pseudoalteromonas sp. 129-1; halotolerant alkaliphilic Bacillus sp. NPST-AK1

Laundry (additive in laundry detergent) and pharmaceutical industries, used as nontoxic antibiofilm agent

Venugopal and Saramma (2007), Wu, Liu, Zhang, Li, and Sun (2015), and Ibrahim, Al-Salamah, El-Badawi, El-Tayeb, and Antranikian (2015)

Ectoine synthase

Sphingopyxis alaskensis

Ectoine biosynthesis [ectoine is a superior skin moisturizer, prevents aging, protects from UV irradiation and skin drying]

Graf, Anzali, Buenger, Pfluecker, and Driller (2008) and Kobus, Widderich, Hoeppner, Bremer, and Smits (2015)

Wax ester synthase (WS), diacylglycerol acyltransferase enzymes (DGAT)

Strains affiliated to OM60 Synthesis and accumulation of neutral lipids group belonging to γ-proteobacteria

Lanfranconi, Alvarez, and Alvarez (2015)

Cold-active and salt-tolerant α-amylase (AmyZ), xylanase/coldactive endo-β-1,4-xylanase

Zunongwangia profunda, Glaciecola mesophila KMM 241

Qin, Huang, and Liu (2013), Liu, Huang, Zhang, Shao, and Liu (2014), Guo, Chen, Sun, Zhou, and Zhang (2009), and Bozoglu, Hundur, Alaylar, Karadayi, and Gulluce (2015)

Applied biology, food industry, and bioethanol production from marine seaweeds; xylo-oligosaccharides production

Continued

Table 2 Details of Marine Enzymes Derived From Bacteria—cont’d Enzyme Enzyme Source

Enzyme Function and/or Applications Reference

Endoglucanase (Cel9P)

Paenibacillus sp. BME-14

Cold-adapted phosphinothricin N-acetyltransferase

Rhodococcus sp. strain YM12 Herbicide detoxification by transgenic crops

Wu et al. (2014)

Serine hydroxymethyltransferase/ hydroxymethyltransferase; β-glucosidase

Alcanivorax sp.; Arthrobacter Enzymatic synthesis of L-serine, nicotianae; Martelella potential for industrial applications mediterranea due to its high enzymatic conversion rate

Yuan, Jiang, Chen, Guo, and Liu (2014), Jiang et al. (2014), and Mao, Hong, Shao, Zhao, and Liu (2010)

L-Glutaminase

Halophilic bacterium— Vibrio costicola

Prabhu and Chandrasekaran (1997)

β-Galactosidase, laminarinase, phosphatase, aminopeptidase

Cyanobacterium synechocystis Confers high tolerance to organism Abdulaziz et al. (2016) spp. to heavy metals like cadmium, lead, mercury, copper, nickel, and zinc with implications in bioaccumulation

Cellulase activity, cold-active mechanism, and industry applications

Anticancer properties

Fu et al. (2009)

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91

grains, shells, sponges, algae, mollusks, corals, plants, fish, etc. (Blackwell, 2011; Bonugli-Santos et al., 2015), and are supposedly the key players in marine habitat. Despite a significant role played especially by the endophytic marine fungi with respect to the bioactive secondary metabolite/compound production (such as terpenoids, steroids, quinones, phenols, and coumarins) possessing antioxidant, antiviral, antibacterial, anticancer, antidiabetic, antifungal, antiprotozoal, antituberculosis, antiinflammatory activities, insecticidal properties, and applications in pharmaceutical and agrochemical industry, these entities occupying the oceans remain under-explored (reviewed by Hamed et al., 2015; Li et al., 2014). Marine fungi secrete several enzymes (Table 3; xylanases, lignin peroxidases, manganese peroxidases, and laccases) that also cause breakdown of complex compounds such as industrial toxins and crude oil components (Atalla et al., 2010). Extremophilic microorganisms produce alkaliphilic enzymes (proteases, cellulases, lipases, and pullulanases) which have tremendous industrial applications (Horikoshi, 1999). Proteases are enzymes with application in detergent formulation industry and are of commercial significance. Proteases and lipases also find their applications in dairy industry. Xylanases are majorly produced by fungi and find prominence in fields of food, feed, beverage, and textile industries and in waste treatment. Another group of enzymes with diverse applications are cellulolytic enzymes required in sugar and ethanol fermentation, detergents, chemicals, pulp and paper, textile industry, animal feed, and food industry (Moubasher et al., 2016). Large-scale production of these enzymes requires optimized culture conditions, i.e., bioprocessing in bioreactors (viz., solid-state fermentation) especially for enzymes such as proteases, chitinases, agarases, peroxidases, glucoamylases, superoxide dismutases, lignin peroxidases, chitinases, and glutaminases (Sarkar et al., 2010). Further, the enzyme needs to be concentrated, isolated, purified using sophisticated and sequential biochemical and biophysical techniques such as ammonium per sulfate precipitation, dialysis, ultracentrifugation, ion-exchange chromatography, acetone precipitation, gel and tangential flow filtration, extraction, hydrophobic interaction chromatography, rechromatography, and speed vacuum concentration. Basically, the aim of employing such purification strategies is to obtain maximum yield of the purest form of the enzyme with continuous product recovery, inexpensive in terms of large-scale and continuous-type production, and maintaining structural conformation of enzyme to retain its specificity and catalytic activity optimum (Bonugli-Santos et al., 2015).

Table 3 Details of Marine Enzymes Derived From Fungus Enzyme Enzyme Source

Enzyme Function and/or Application

Reference

Niyonzima and More (2014)

Alkaline protease

Scopulariopsis spp.

Useful in detergent formulations

Alginate lyase, amylase, cellulase, chitinase, fructosyl-aminooxidase, fucoidanase, glucanase, galactosidase, glucosidase, glucosaminidase, hexoseaminidase, inulinase, keratinase, ligninase, lipase, nuclease, phytase, polygalacturonase, protease, speroxide dismutase, and xylanase

Pestalotiopsis sp., Aureobasidium pullulans N13d, Penicillium janthinellum P9, Debaryomyces hansenii C-11, Aspergillus oryzae, Penicillium canescens, Trichoderma aureviride KMM4630, Aspergillus awamori BTMFW032, Penicillium melinii, Flavodon flavus, etc.

Bonugli-Santos et al. (2015) Implicated in Fungal Marine Biotechnology (Biotechnology— hydrolytic and oxidative enzymes; Environmental Biotechnology—enzymes degrading textile effluents and polycyclic hydrocarbons; and Industrial Biotechnology—enzymes related to manufacture of chemical, fuel, food [dairy, baking], beverage, agriculture related, textiles, cosmetics, etc.)

Amylase, cellulase, chitinase, gelatinase, lipase

14 Fungal genera: Penicillium, Significant role in remineralization and several industrial applications Aspergillus, Scopulariopsis, Cephalosporium, Humicola, Gymnoascus, Endomysis, Zygorhynchus, Trichoderma, Zalerion, Pleospora, Chaetomium, Phoma, Botryphialophora, unidentified

Smitha, Correya, and Philip (2014)

Laccase, lipase, cellulase, peroxidase, manganese peroxidase

Nigrospora species and Arthopyrenia species—marine sponge Trematosphaeria mangrovei

Baldrian (2006), Atalla et al. (2010), Atalla, Zeinab, Eman, Amani, Abd, and AtyAbeer (2013), Li, Singh, Liu, Pan, and Wang (2014), and Passarini, Ottoni, Santos, Lima, and Sette (2015)

Cause breakdown of several environmental compounds, such as industrial toxins and crude oil components; active role in ecological cycles of coastal ecosystem, significance in bioremediation

L-Glutaminase

Marine Beauveria sp.

Uncoupling protein 1 (UCP1)

U. pinnatifida, Hijikia fusiformis, Antiobesity enzyme and Sargassum fulvellum

Tannase (tannin acyl hydrolase) Aspergillus awamori BTMFW032

Cellulase, xylanase, and pectinase

Alternaria alternata, Aspergillus terreus, Cladosporium cladosporioides, Emericella nidulans, Fusarium solani, Cochliobolus australiensis

Anticancer properties

Sabu, Keerthi, Kumar, and Chandrasekaran (2000) € € and Hamed, Ozogul, Ozogul, Regenstein (2015)

Hydrolysis of ester and depside bonds to Beena (2010) synthesize gallic acid and glucose; gallic acid is a substrate for production of antibacterial drug trimethoprim, synthesis of propyl gallate, an antioxidant used in food industry; and catechin gallates are used in manufacture of instant tea, coffeeflavored soft drinks, flavor improvement in grape wine, beer, and fruit juice clarification, to enhance antioxidant activity of green tea, cleavage of polyphenolics, determination of structure of naturally occurring gallic acid esters Implicated in food, feed, beverage, textile Moubasher, Ismail, Hussein, and Gouda (2016) industries and in waste treatment; fermentation of sugars and ethanol; for production of detergents, chemicals, pulp and paper; required in textile industry, animal feed, and food industry

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4.4 Sponges Sponges (phylum Porifera) are aquatic invertebrates known to inhabit the earth since 700–800 million years with about 15,000 known species. They easily thrive in temperate, salty water, deep oceans, or coastal area and have attracted attention of researchers worldwide due to their rich bioactive secondary metabolites (namely amino acids, aliphatic cyclic peroxides and sterols, macrolides, nucleosides, porphyrins, terpenoids) and enzymes. These components possess medicinal and pharmacological relevance due to antibacterial, anticoagulant, antifungal, antimalarial, antituberculosis, antiviral, antiinflammatory, immunosuppressive, neurosuppressor, and antitumor activities and have been successfully harnessed for several industrial and biotechnological research, drug discovery, and commercial applications which have reached clinical trials phases I, II, or even to the market (reviewed by Lira et al., 2011; Thakur & Muller, 2004; Thomas, Kavlekar, & Lokabharathi, 2010). Symbiotic association of bacteria and sponge is essentially observed between bacteria from aquatic habitat, its own intracellular microbiota, or those dwelling within the sponge mesophyll. Similarly, ubiquitous association of planktonic and benthic diatoms, fungi, unicellular algae, archaea, heterotrophic bacteria, cyanobacteria, facultative anaerobes, etc., is also observed (Wang, 2006). Among several bioactive metabolites, enzymes produced by (especially halichondrid) sponge-associated microbes are gaining importance due to their therapeutic potential and others due to high profile of applications they offer (Table 4). Out of 10 bacterial phyla involved in symbiotic associations (Thomas et al., 2010), the polyketide synthase (PKA) genes found in Actinobacteria, Bacillus, Sulfitobacter, and Pseudovibrio and PLA2 (Phospholipase A2) which are crucial precursors for synthesis of secondary metabolites serve as host defense mechanism for marine sponges with bacterial symbiotic associations due to their antibacterial properties [e.g., Streptomyces dendra sp. nov. MSI051 isolated from marine sponge Dendrillanigra possess antibacterial activity found in west coast of India] (Piel et al., 2004; Selvin, 2009; Selvin & Lipton, 2004). Chitinase isolated from the marine Streptomyces sp. DA11 found in association with South China sponge Craniella australiensis possesses antifungal activity. In addition, marine-derived chitinase has higher pH and salinity tolerance, thus making them attractive candidates for several biotechnological applications (Han, Yang, Zhang, Miao, & Li, 2008). Marine sponges are emerging as promising sources for bone tissue engineering because they possess a characteristic porous architecture simulating a

Table 4 Details of Marine Enzymes Derived From Sponge Enzyme Enzyme Source

Enzyme Function and/or Applications

Reference

α-Carbonic anhydrases

Calcareous sponge—Sycon ciliatum and Biomineralization enzyme required in sponge Voigt, Adamski, Sluzek, Leucosolenia complicata spicule formation and Adamska (2014)

Aeroplysinin 1-specific nitrile hydratase

Sponge (Aplysina cavernicola)-associated Implicated in converting aeroplysinin-1 into Lipowicz, Hanekop, bacteria dienone amide verongiaquinol which is used Schmitt, and Proksch as a defensive compound (2013)

Carboxymethylcellulase, Sponge (viz. Spirastrella sp., Phyllospongia Novel source for industrial, commercial sp., Ircinia sp., Aaptos sp., Azorica sp., and applications, useful for cycling of organic protease, matter in marine environments Axinella sp.) deoxyribonuclease, gelatinase, lipase, phosphatase, and urease Extracellular hydrolytic enzymes-(α)-amylase, (alkaline) phosphatase

Marine sponge Dysidea granulosa, Sigmadocia fibulata-associated bacteria Gammaproteobacteria, Vibrionales— chief source of multiple enzyme production; salt-tolerant Actinomycete Streptomyces sp. from marine sponge Ircinia sp.; Callyspongiidae

Wide applications in starch processing, food, paper, detergent, fermentation, textile, and pharmaceutical industry; alkaline phosphatase expression in marine sponge implies its osteogenic potential, and thus it may serve as potential new scaffold for bone tissue engineering

Mohapatra, Bapuji, and Sree (2003)

Feby and Nair (2010), de Souza and Magalha˜es (2010), Lin (2011), and Krishnakumar, Bai, and Premkumar (2015)

Continued

Table 4 Details of Marine Enzymes Derived From Sponge—cont’d Enzyme Enzyme Source

Enzyme Function and/or Applications

Reference

Helps to maintain constant ATP levels during Harcet, Perina, and Plesˇe (2013) exercise and hypoxic condition, and thus maintains redox balance in cytosol during glycolysis under anoxic conditions

Opine dehydrogenases

Marine sponge—Suberites domuncula

Phospholipase A2 (PLA2)

Sponge-associated bacterium belonging Implicated in conferring defense potential to Shen and Cho (1995), Selvin, Ninawe, Kiran, to phylum Porifera the host sponge during its symbiotic and Lipton (2010), and association with bacteria; useful in food, cosmetics, pharmaceutical industry, Kishimura (2012) implicated in hypercholesterolemia therapy

Silicatein

Marine sponge—Tethya aurantium, Suberites domuncula

Cha et al. (1999) and Implicated in biosilica formation, silica metabolism, production of bone replacement Wang et al. (2011) material and its potential in the treatment of osteoporosis

Biotechnological Applications of Marine Enzymes

97

bone scaffold, comprising spongin, analogous to vertebral collagen, a natural polymer essential for tissue regeneration, possess osteogenic properties, due to biosilica, other minerals, compounds, etc., that support cell growth and stimulate mineralization and bone formation in in vitro models such as human multipotent stromal cells, mouse primary osteoblasts, cell lines, and bone marrow cells (reviewed in Granito, Custo´dio, & Renno´, 2016). Biosilicification is a process exclusive to marine organisms inclusive of sponges (10,000 species). Sponges (e.g., Tethya aurantium, Petrosia ficiformis, Suberites domuncula) consist of cathepsin L-related enzyme, termed (α, β)silicatein (Cha et al., 1999; Shimizu, Cha, Stucky, & Morse, 1998) and later versions of biocatalytically active recombinant silicatein catalyze silica formation from monomeric silica precursors like synthetic tetraethoxysilane, whereas silicase (Schr€ oder et al., 2003) is a silica-degrading/metabolizing enzyme required for spicule formation–biomineralization and has revolutionized fabrication process in nanobiotechnology and biomedicine. Marine silicatein-mediated silica glass production at low temperatures, pH, and near neutral pH has implications in biomimetic fabrication for nanostructured materials and devices in opto- and microelectronics industry (reviewed in Schr€ oder, Wang, Tremel, Ushijima, & M€ uller, 2008). Several hydrolytic enzymes such as α-amylase, carboxymethylcellulase, protease, urease, lipase (alkaline) phosphatase, heat-tolerant acetylcholinesterase have assumed prominent place in the production of beverages, food, confectionery, textiles and leather processing, waste water treatment, for starch processing, fermentation, paper, detergents, and most importantly manufacture of pharmaceuticals (Mohapatra et al., 2003, Wang, 2006). Role of mutual symbiotic mechanism between host and its associated microbe and resultant secondary metabolite production with pharmaceutical and medicinal relevance are poorly understood. To obtain these and further refine this system for scale-up production and efficient recovery of final products are the reigning topics of interest among marine researchers. Besides, in vitro ability of such microbes to synthesize metabolites with efficiency similar to in vivo scenario for over several generations may be a serious bottleneck calling for further research. Researchers wish to transfer symbiont biosynthetic genes into cultivable microbes for further biotechnological purposes. PKS gene-based molecular techniques may help in screening of pharmaceutically valuable strains and to predict related compounds. Discovery of new genes, enzymes, and bioactives using the latest sophisticated molecular genetic techniques (viz., metagenomics, etc.) and refining culture techniques mimicking in vivo state may boost production of marine biocatalysts

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S. Parte et al.

and drugs manifold. Identification of novel candidate biosynthetic genes with therapeutic significance may be the next avenues which are already being pursued. Fisch et al. (2009) devised a strategy based on chemical moieties to quickly access biosynthetic gene clusters for PKA gene from marine sponge–bacterial association of Psammociniaaff bulbosa and discovered complete pathway for a rare and potent antitumor agent psymberin related to pederin family.

5. CHALLENGES ENCOUNTERED IN HARNESSING MARINE RESOURCES In order to harness the tremendously valuable marine resources, access to these marine microbes and several tools of manipulation is requisite. Due to the harsh and extreme conditions, oceanic habitat itself poses a challenging environment (benthic sea beds, deep-sea vents, permafrost regions, or pelagic zones). Hence, these vast resources largely remain untouched/ untapped, while access only to intertidal and shallow subtidal environment is possible. Standard culturing methods for marine microbes face several constraints; as a result, almost 99% of marine species remain uncultured and hence unexplored. In order to explore marine biodiversity in deeper oceanic habitat, sophisticated and expensive equipment such as submersibles, remote-operated vehicles, automated sampling devices, to reach and collect samples from inaccessible places is required (Cheung, Watson, & Pauly, 2013). Further global warming and climatic changes impose detrimental effect on the oceanic ecosystem and the vast array of resources that they offer (Thakur & Thakur, 2006). Biocatalysts produced naturally by marine microbes may be present in scarce concentration are mixed up with many other enzymes, or their yield may be low or may exist in trace amounts. Mass production entails cultivation of the appropriate bacteria in bulk which could be marred by an undesirable by-product or lethal toxin formation and possibility of crosscontamination (Gurung, Ray, Bose, & Rai, 2013). Variation depending upon enzyme used, its application, carrier system used, designing performance of the immobilized enzyme for a particular bioprocess and thus predicting its performance/efficiency in advance is a major hurdle (Cao, 2005). Inability of retrieving a “sustained and reliable” harvest of marine organisms yielding marine enzyme, insufficient quantities of marine harvest for study completion, and difficulties while culturing marine organisms (Bhadury, Mohammad, & Wright, 2006) are other challenges. Several

Biotechnological Applications of Marine Enzymes

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factors may interfere with the utility of a specific enzyme especially for therapeutic purposes (i) due to its molecular size which may hinder its incorporation within the cells; (ii) due to host cell immune response within host organism/cell postintroduction of a foreign enzyme protein; and (iii) to maintain high degree of purity and specificity (Nigam, 2013). It becomes imperative to utilize these resources prudently without adversely affecting their natural abundance. Besides the conventional methods of mariculture/aquaculture, fermentation for large-scale cultures combined with overexpression of the enzyme in a heterologous organism allowing optimal enzyme activity is required. State of the art of bioreactor engineering and bioprocess designing for cultivating suspended organisms or immobilized cells for production of marine enzymes require to be adopted systematically (Sarkar et al., 2010). By employing cutting-edge and nonconventional multidisciplinary technologies, some of the challenges would be overcome.

6. FUTURE PROSPECTS Development of target-specific and cost-effective enzyme technology is a need. Several revolutionary modifications and improvisations to currently existing procedures are required. For instance, complete purification of pure enzymes from marine organisms, determination and characterization of their active sites, cross-linking of enzymes for stabilization, and enzyme immobilization need improvement. Multienzyme immobilization with colocalization on nanoparticles, in silico approaches for protein engineering, de novo assembly of contig mapping, next-generation pyrosequencing, etc., to discover novel pathways, creation of transcriptome and metabolome database for future biosynthetic engineering in order to establish improved strains and ensure transfer of desirable traits are stepping stones toward establishing efficient and cost-effective “Marine” Enzyme Technology. Other approaches include preservation of rare resources obtained from deep oceans, derivation of cell lines, and their long-term cryopreservation, gene cloning, sequencing, use of recombinant microbes for improving enzyme stability by site-directed mutagenesis, protein engineering, study of crystallography of the enzymes, etc., thus enhancing further potential of marine enzymes for bioprocess technology, biotransformation, bioremediation, biosensor development, etc., with greater efficiency than those used currently. A detailed understanding of the structure–function relationship is the most crucial aspect for commercialization of any biocatalyst for further high-throughput biotechnological applications.

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CHAPTER FIVE

Biomedical Applications of Enzymes From Marine Actinobacteria K. Kamala1, P. Sivaperumal Center for Environmental Nuclear Research, Directorate of Research, SRM University, Kattankulathur, India 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Marine Actinobacteria and Their Enzymes 2.1 L-Asparaginase 2.2 Cholesterol Oxidase 2.3 Laccase 2.4 Other Enzymes 3. Biological Activities of Enzymes From Marine Actinobacteria 4. Pharmacological Activity of Marine Organism–Associated Actinobacteria 5. Conclusion Acknowledgments References

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Abstract Marine microbial enzyme technologies have progressed significantly in the last few decades for different applications. Among the various microorganisms, marine actinobacterial enzymes have significant active properties, which could allow them to be biocatalysts with tremendous bioactive metabolites. Moreover, marine actinobacteria have been considered as biofactories, since their enzymes fulfill biomedical and industrial needs. In this chapter, the marine actinobacteria and their enzymes’ uses in biological activities and biomedical applications are described.

1. INTRODUCTION Over the last three decades, studies on enzyme technology and enzymology have progressed significantly, and many industrial applications from enzymes are used in the native or immobilized stage. Their applications Advances in Food and Nutrition Research, Volume 80 ISSN 1043-4526 http://dx.doi.org/10.1016/bs.afnr.2016.11.002

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2017 Elsevier Inc. All rights reserved.

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include materials processing, textiles and food industries, detergents, biochemical and biotechnological purposes, chemical industry, and pharmaceutical uses (Cetinus & Oztop, 2003). These enzymes are dynamic to living organisms, particularly regulating different biochemical events such as catabolism, metabolism, cellular signal transduction, cell cycling, and development. However, enzymes are frequently related to human disease, and they are also supported by the analysis of disease using molecular techniques. In humans, many disorders are caused by the dysfunction of enzymes, as well as overexpression or hyperactivation of enzymes (Otto & Schirmeister, 1997). Since the mid-1950s, there has been a significant increase in both the measurement of enzyme activities and the use of purified enzymes obtained from marine sources for different medical practices. Recently, various kinds of marine enzymes have been isolated, purified, and used for clinical trials. Further developments in enzyme production have made available a number of potential enzymes, which have been used in enzyme therapy. Till now, the researchers are interest in learning more about a largely unexplored monarchy of marine enzymes for their biological activity purposes. In particular, a marine enzyme from microorganisms has a particular protein molecule that is not found in any terrestrial organisms. Besides marine microorganisms of bacteria, actinobacteria, fungi, and many other marine organisms such as plants, algae, prawns, crabs, snakes, and fishes have been studied to tap the resource of the marine world. Specifically, marine actinobacteria are the biotechnologically and most economically precious prokaryotes group. They are abundant producers of antibiotics and significant suppliers to the pharmaceutical industry and could produce an extensive variety of secondary metabolites such as enzymes, enzyme inhibitors, antibiotics, pigments, and immunosuppressive agents (Manivasagan, Venkatesan, Sivakumar, & Kim, 2013). Therefore, studies on marine enzymes and their potential biological activities are more important. In this context, this chapter gives detailed information on enzymes from marine actinobacteria and their applications in biological activities and usage in modern technology.

2. MARINE ACTINOBACTERIA AND THEIR ENZYMES Earlier, actinobacteria were universally known as Actinomycetes and the term Actinomycetes belongs to the Actinomycetales order, which is the main subdivision of the prokaryotes kingdom. Later, Actinomycetales were

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identified as higher bacteria possessing transitional properties between bacteria and fungi (Suneetha & Khan, 2010). In addition, actinobacteria are Gram-positive microorganisms that tend to grow slowly as branching filaments, and they look like fungi, as their mycelial colony growth is in filamentous forms that may be aerial alone or both substrate and aerial. The substrate mycelium was stretchable with a media, and the spore bearing hyphae of aerial mycelium have a longer length. Approximately 240 actinobacterial genera identified and they fall into 1 of 5 subclasses, 9 orders, and 55 families, embracing 3000 species (Goodfellow & Fiedler, 2010). Recently, researchers found the existence of marine actinobacteria and their wide distribution in different marine ecosystems (Ghosh, Barua, & Chakraborty, 2011; Haritha, Kumar, Mohan, & Ramana, 2010; Sathiyaseelan & Stella, 2011), since the marine environment has different types of physicochemical parameters such as pressure, salinity, temperature, light, and nutrients. The marine actinobacteria have adapted themselves to survive in the marine environment and they require Na+ for their growth and to maintain the osmotic pressure environment to protect their cellular integrity (Goodfellow & Williams, 1983). They have unique metabolic and physiological processes, which ensure their survival in extreme habitats and also offer them the possibility of producing compounds with many interesting pharmacological activities (Maskey et al., 2004). In addition, marine microorganisms have been obtainable from sediment and water samples by biotechnologists in and around onshore and deep sea environments. Therefore, reports of marine microbial enzymes from these sources have been numerous in past decades. Especially, marine actinobacteria are recognized to produce various enzymes such as protease, amylase, cellulase, and lipase, which have enormous biomedical applications. Still, the usage of enzymes is very limited in the medical/pharmaceutical and food-processing industries. Potential enzymes from marine microorganisms have been reported by several researchers, and some of these enzymes are given in detail.

2.1 L-Asparaginase marine actinobacteria Streptomyces spp. was isolated from sediment samples taken from the southwest coast of India, and these isolates showed 64.07 IU/mg as a maximum specific activity at pH 8 in 60–80°C. These enzymes have 140-kDa molecular weights, and they also showed the cytotoxic effects on acute T-cell leukemia and chronic myelogenous leukemia (Dhevagi & Poorani, 2006). Similarly, Basha, Rekha, L-Asparaginase-producing

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Komala, and Ruby (2009) explored the production of extracellular antileukemic enzyme L-asparaginase from microorganisms using low-cost substrate soybean meal under solid-state fermentation and their enzyme production was also observed in the range between 24.6 and 49.2 IU/mL. Later, Amena, Vishalakshi, Prabhakar, Dayanad, and Lingappa (2010) also isolated the novel L-asparaginase-producing Streptomyces gulbargensis and their purification (82.12-fold), and observed the molecular weight (85 kDa), and stable alkaline (55%). Also, 17 L-asparaginase-producing Streptomyces were isolated from Gulf of Mannar marine sediment samples using starch casein agar. Out of those 17, 5 strains were L-asparaginase positive (Jayam & Kannan, 2012). A total of three L-asparaginase-producing marine actinomycetes were isolated from marine sediments of the southern India coastal region. These isolates were screened by semiquantitative plate assays, which showed 1.92, 1.48, and 1.46 U/mL specific enzyme activities (Selvam & Vishnupriya, 2013). Similarly, Krishnamayi, Poda, and Vijayalakshmi (2013) also isolated the L-asparaginase-producing potential marine actinobacteria from the nizampatnam mangrove ecosystem, and their maximum specific activity was 7.42 IU/mL at pH 8 in 30°C. Meena et al. (2015) reported the L-asparaginase enzyme from Streptomyces griseus NIOT-VKMA29, and their enzyme production also greatly increased to 123 IU/mL by gene (ans-A) expression in Escherichia coli M15. Recently, Dhevagi and Poorani (2016) also reported the L-asparaginase-producing actinobacterial isolate of DS8 from the Tuticorin coastal area, which showed 45.18 IU/mg of specific activity. Mangamuri, Muvva, and Poda (2016) observed the extracellular L-asparaginase enzyme from Pseudonocardia endophytica VUK-10, which was isolated from the mangrove environment. The enzyme was 96-fold and showed specific activity of 702.04 IU/mg with a 61% yield, and the molecular weight was 120 kDa.

2.2 Cholesterol Oxidase Generally, actinobacteria produces a high level of extracellular cholesterol oxidase (CO) enzyme (Yazdi, Zahraei, Aghaepour, & Kamranpour, 2001). Ivshina, Grishko, Nogovitsina, Kukina, and Tolstikov (2005) reported the CO enzyme-producing Rhodococcus sp. and it is used to measure the cholesterol in biological fluid samples of the human mammary gland (Donova, 2007). An important actinobacterial genus of Mycobacterium sp. could produce CO enzymes (Wilmanska et al., 1995), as well as some other genera such as Rhodococcus sp. (Fernandez et al., 2011), Nocardia sp.

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(Horinouchi, Ishizuka, & Beppu, 1991), and Brevibacterium sp. (Fujishiro et al., 1990), and Streptomyces sp. (Ishizaki, Hirayama, Shinkawa, Nimi, & Murooka, 1989) also was found to produce CO. In addition, intracellular CO-producing Arthrobacter sp. (Doukyu, 2009), Corynebacterium sp. (Rhee, Kim, & Park, 2002), Nocardia sp. (Doukyu, 2009), and Streptomyces spp. (Kimberley, Ignatius, Sampson, & Vrielink, 1999; Nishiya et al., 1997) were isolated from various sources, and their molecular weight ranges were observed to be between 35 and 70 kDa at pH values ranging from 6 to 8. Similarly, extracellular CO-producing Brevibacterium sp. (Croteau & Vrielink, 1996), Rhodococcus equi (Bokoch, Devadaoo, Palencsar, & Burgess, 2004), and Streptomyces spp. (Sampson & Vrielink, 2003) were reported, and their molecular weights ranged from 30 to 61 kDa. Later, Niwas, Singh, Singh, Tripathi, and Tripathi (2013) isolated the CO from Streptomyces sp. with different supplementations of maltose, lactose, sucrose, peptone, soyabean meal, and yeast extract to enhance enzyme production, and the maximum production was noted in the basal production medium as 14.3-fold at optimum pH 7 in 37°C with a molecular weight of 62 kDa.

2.3 Laccase Laccase enzymes from S. griseus and Streptomyces lavendulae were isolated and molecular weights of 114 and 73 kDa, respectively, were determined (Suzuki et al., 2003). Similarly, Arias et al. (2003) reported that the laccase enzyme from the actinobacteria of S. cyaneus has a molecular weight of 75 kDa and that the highest production of enzyme was observed in pH 4.5 at 70°C. Another actinobacterial species of S. coelicolor and S. psammoticus produced the laccase enzyme in pH 9.4 at 60°C and pH 8.5 at 45°C, respectively (Niladevi, Jacob, & Prema, 2008), and their molecular weight was measured (69 and 43 kDa, respectively). Later, Gunne and Urlacher (2012) and Lu et al. (2013) also observed the laccaseproducing enzyme from Streptomyces ipomoea and Streptomyces sp., which has 79- and 38-kDa molecular weights, respectively, at high temperature (40° C). Thermobifida fusca BCRC 19214 (Chen, Hsieh, Cheepudom, Yang, & Meng, 2013) was found to produce thermoalkali-stable laccase used in hair color applications.

2.4 Other Enzymes The hydrolytic enzyme xylanase was reported from marine actinobacteria strain of Thermomonospora fusca (Bachmann & McCarthy, 1989) and

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Streptomyces sp. Ab106 (Elegir, Sykes, & Jeffries, 1995) at 60°C and pH 7 and 9, respectively. Likewise, Streptomyces thermoviolaceus OpC-520 (Tsujibo et al., 1992), Streptomyces viridosporus T7A (Magnuson & Crawford, 1997), and Thermomonospora curvata (Stutzenberger & Bodine, 2008) also produced alkaline and thermostable xylanase enzymes. In addition, Sathyapriya, Stalin, and Selvam (2012) isolated the xylanase-producing Streptomyces albus MAC6 strain from marine sediment samples from the south coast of India. The maximum production of xylanase from S. albus was 302.50  0.36 IU/mL at pH 10 and 50°C with 10% of substrate concentration. Similarly, the xylanaseproducing marine actinobacteria Streptomyces sp. was isolated from mangrove sediment samples from the Pichavaram mangrove environment, and these strains synthesized the maximum amount of enzyme in 10 days at 35°C and pH 6.8 (Umadevi & Ragunathan, 2013). Liu, Zhao, and Bai (2013) demonstrated that ribosome engineering can be successfully applied to improve enzyme production in Streptomyces sp. M11, which was isolated from marine sediment samples from Xiaoping Island in Dalian, China. A novel alkali-tolerant and thermostable inulinase-producing marine actinobacteria, Nocardiopsis sp. DN-K15,was isolated from the sediments of Jiaozhou Bay in China. This Nocardiopsis sp. DN-K15 strain produced 25.1 U/mL of inulinase with 60 h of fermentation time at pH 8 and 60°C (Lu, Li, & Guo, 2014). Similarly, inulin-hydrolyzing Streptomyces sp. KF5 (Reddy, Deepthi, Parameshwar, & Sulochana, 2015) and Streptomyces sp. CP01 (Laowklom, Chantanaphan, & Pinphanichakarn, 2012) were isolated, and these strains were produced 1.60 U/mL of inulinase at 28°C and pH 8. Few reports exist about bifunctional thermostable alginate lyase from Isoptericola halotolerans CGMCC 5336 (Dou et al., 2013), and it is used as an anti-inflammatory agent, antitumor, and anticoagulant agent (Iwamoto et al., 2005). The actinobacterial strain Thermobifida fusca also produced acetyl xylan esterase at 80°C and pH 8 (Yang & Liu, 2008). Marine actinobacterial strains Rhodococcus rhodochrous, R. erythropolis, Corynebacterium spp., Brevibacterium spp., Streptomyces spp., and Mycobacterium spp. are known to produce carboxyl oxidase enzyme (MacLachlan, Wotherspoon, Ansell, & Brooks, 2000), which is used in the conversion of cholesterol (Pollegioni, Piubelli, & Molla, 2009), and carboxyl oxidase catalyzes cholesterol to Cholest-4-en-3-one by oxygen reduction at C3 to hydrogen peroxide (Kreit & Sampson, 2009). In addition, Rhodococcus sp. and Streptomyces spp. were producing CO, and the strains were also identified as CO-producing organisms (Kumari & Kanwar, 2012). Moreover, alkali thermo stable dextranase was isolated from Streptomyces sp. NK458

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(Purushe, Prakash, Nawani, Dhakephalkar, & Kapadnis, 2012). Recently, Martinez, Wu, Sanishvili, Liu, and Holz (2014) reported on thermostable nitrile hydratase from the actinobacteria of Pseudonocardia thermophila.

3. BIOLOGICAL ACTIVITIES OF ENZYMES FROM MARINE ACTINOBACTERIA Approximately 23,000 antibiotics have been reported to be produced from microorganisms. Among them, 10,000 antibiotics have been isolated from actinobacteria. Of these, the Streptomyces genus mainly has the ability to produce a wide variety of primary and secondary metabolites having bioactive potential, including antibiotics. In addition, the actinobacterial group has numerous biosynthetic activities, which are absent in other microbial groups. This enormous diversity, besides its less utilization, scientists have been interested in exploring potential metabolites from them. The Streptomyces genus (with 500 species) is signified in nature by the major species among all other genera of actinobacteria. These actinobacterial enzymes in the form of any functional groups are involved in biomedical applications, whether directly or indirectly. One of the monofunctional or bifunctional enzymes called polyketides is a huge and structurally diverse family of natural products. Few of the polyketides with medical importance, such as epirubicin, doramectin, and compounds like aurantimycin, kirromycin, chartreusin, polyketomycin, concanamycin, and lysolipin, have been used yet for therapeutic treatments (Weber, Welzel, Pelzer, Vente, & Wohlleben, 2003). Cytotoxic polyketides of Daryamides A, B, and C were isolated from the marine Streptomyces CNQ-085 strain, and Daryamides B and C showed weak-to-reasonable cytotoxicity against the human colon carcinoma cell line of HCT-116. However, Daryamide A showed significantly more potent cancer cell cytotoxicity, with an IC50 of 3.15 μg/mL, and less antifungal activity against Candida albicans, which was also shown by polyketides (Asolkar, Jensen, Kauffman, & Fenical, 2006). The new polyketides Actinofuranones A and B were isolated from the marine-derived Streptomyces strain CNQ766, and they exert weak in vitro cytotoxicity activity against mouse macrophages and splenocyte T-cells (Cho et al., 2006). A novel polycyclic polyketide (Abyssomicin C) was isolated from marine Verrucosispora sp. (Riedlinger et al., 2004), and it showed itself as a potential target against para-aminobenzoic acid biosynthesis. In addition, Abyssomicin C showed antibacterial activity against Gram-positive bacteria, including multiple

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resistant clinical isolates and vancomycin-resistant Staphylococcus aureus, and its similarities thus have the potential to be antimicrobial agents against multidrug-resistant pathogens (Rath, Kinast, & Maier, 2005). Higher type I polyketide-derived compounds of Arenicolides were obtained from the Salinispora arenicola strain CNR-005, isolated from the island of Guam, and it has moderate cytotoxicity against the human colon adenocarcinoma cell line HCT-116, with an IC50 value of 30 μg/mL (Williams et al., 2007). In addition, the aromatic polyketide glycoside of Chartreusin also was isolated from Streptomyces chartreusis, and it acts as potent antitumor agent (Xu, Jakobi, Welzel, & Hertweck, 2005). L-Asparagine is a therapeutic enzyme for lymphosarcoma and acute lymphoblastic leukemia (ALL) in humans (Ahmady, Sara, El-Ewasy, & El-Shweihy, 2014; Narta, Kanwar, & Azmi, 2007; Savitri & Azmi, 2003). The commercial product of Erwinase, Leukine, Elspar, and Kidrolase is microbial L-asparaginase used in the treatment of ALL (Verma, Kumar, Kaur, & Anand, 2007). L-Asparaginase creates one of the greatest biomedical and biotechnologically essential groups of beneficial enzymes, accounting for 40% of enzyme sales worldwide. The most common therapeutic indications of L-asparaginase mainly focus on treating children for acute lymphocytic leukemia, as well as other treatments of Hodgkin disease, acute myelomonocytic leukemia, acute myelocytic leukemia, and chronic lymphocytic leukemia, reticlesarcoma, lymphosarcoma, and melanosarcoma (Stecher, de Deus, Polikarpov, & Abrahao-Neto, 1999). In addition, L-asparaginase contributes one-third of the global requirements of antileukemic and antilymhoma agents, far more than other therapeutic enzymes (Hosamani, 2012), and their usage has been more extensive in antileukemia therapy in acute lymphoblastic leukemia disease (Jain, Zaidi, Verma, & Saxena, 2012; Patil, Coutsouvelis, & Spencer, 2011). Glutaminase-free L-asparagine was produced by Streptomyces albidoflavus (Narayana, Kumar, & Vijayalakshmi, 2008), S. gulbargensis (Amena et al., 2010), and S. venezuelae (Saxena, Upadhyay, & Kango, 2015), and it is used as a therapeutic enzyme because glutaminase activity in L-asparaginase may cause side effects (viz., anaphylaxis, stroke, pancreatitis on enzyme therapy). The alkaline phosphatase enzyme was used in clinical diagnoses for kidney damage and toxemia of pregnancy (Liu & Li, 2000). Also, immobilized enzymes are used as monoclonal antibodies because of their substratespecific activity (Delvaux & Demoustier-Champagne, 2003). The laccase enzyme was used in the synthesis of methyl-1,4-hydroquinone and 2,3dimethyl-1,4-hydroquinone, as well as 14 novel cephalosporins, penicillins,

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and carbacephems by amination with lactamase. The authors concluded that this is a promising method to increase the range of antibiotic production, and that the antibiotics are inhibiting the various Gram-positive bacteria (Mikolasch et al., 2016).

4. PHARMACOLOGICAL ACTIVITY OF MARINE ORGANISM–ASSOCIATED ACTINOBACTERIA Multidomain polyketide synthases are producing a structurally diverse group of polyketides compounds. Most of the polyketides compounds that are present in marine microorganisms have relevant biomedical and industrial importance. In addition, the presence of many polyketide genes in actinobacteria (sponge-associated) has been effectively discovered using universal primers (Kim & Fuerst, 2006; Schirmer et al., 2005). Another group of the ansamycin family of antibiotics of rifamycins was used to cure tuberculosis and other microbial infections (Floss & Yu, 2005). Kim, Hewavitharana, Shaw, and Fuerst (2006) also predicted sponge-associated Salinispora’s rifamycin by ketosynthase gene sequence analysis. The anthracyclines are isolated from marine-derived microorganisms and used in cancer chemotherapy (Arcamone & Cassinelli, 1998). Doxorubicin and daunorubicin compounds produced by the marine actinobacteria of S. peucetius have potential to be effective against different types of cancer cells (Newman & Cragg, 2007). The isolation and identification of the novel anthracyclines compound have been attempted for the past two decades (Minotti, Menna, Salvatorelli, Cairo, & Gianni, 2004). Anthracyclines have potential activity against the various arrays of solid tumors and hematological distortions and their medical applications are delayed due to tumor resistance and heavy toxicity to healthy cells. In addition, two new anthracyclines, such as tetracenoquinocin and 5-iminoaranciamycin, were isolated from Haliclona sp. sponge–associated Streptomyces sp. Sp080513GE-26 (Motohashi, Takagi, & Shinya, 2010), and the tetracenoquinocin compound showed only feeble cytotoxic activity against HL-60 and HeLa cancer cells. A group of aromatic polyketides (Angucyclines) also was isolated from marine actinobacteria, which exhibit enzyme inhibitory, antibacterial, antiviral, and antitumor activities (Metsa-Ketela, Palmu, Kunnari, Ylihonko, & Mantsala, 2003; Rohr et al., 1993). The marine actinobacteria of Saccharothrix espanaensis were isolated from the Anadara broughtoni mollusk samples collected from Peter the Great Bay, in the Sea of Japan, Russia,

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and it produced angucyclines, saccharothrixins A–C compounds, which have potential antimicrobial activities (Kalinovskaya et al., 2010). Moreover, a new benza anthracene derivative compound of Mayamycin was isolated from the marine Streptomyces sp. which was associated with the Halichondria panacea sponge. These Mayamycin compound showed potential inhibitory activity against many clinical bacterial and other relevant strains, such as Pseudomonas aeruginosa and methicillin-resistant S. aureus, human skin disease–related bacteria of Dermabacter hominis and Brevibacterium epidermidis, Propionibacterium acnes, and phytopathogenic bacterium of Xanthomonas campestris. Furthermore, it showed cytotoxic effects in the mouse fibroblast cell line NIH-3T3 (Schneemann, Kajahn, et al., 2010; Schneemann, Ohlendorf, et al., 2010). The γ-pyrones are flexible intermediates in organic chemistry for biologically important molecule production (Lin et al., 1995). A total of 13 structurally related nocapyrones within the members of γ-pyrone were isolated (Schneemann, Kajahn, et al. 2010; Schneemann, Ohlendorf, et al., 2010; Lin et al., 2013). In addition, four new γ-pyrones, nocapyrones A–D, were attained from marine actinobacteria of Nocardiopsis sp. HB383, which was associated with bioactive marine sponge Halichondria panicea. Among them, nocapyrones A and B showed antibacterial potential against Staphylococcus lentus and Bacillus subtilis (Schneemann, Kajahn, et al., 2010; Schneemann, Ohlendorf, et al., 2010). Another three nocapyrones A, B, and C were isolated from cone snail-associated marine Nocardiopsis alba CR167 (Lin et al., 2013) and nocapyrones B showed potential activity against nearly all neuronal cell types. Unidentified ascidian-associated Streptomyces sp. YM14-060 produced the piericidins C7 and C8 compounds (Hayakawa et al., 2007) and these piericidins show cytotoxicity against RG-E1A-7 cells; and without any cytotoxic cell death, these compounds inhibited the growth of neuro-2a cells. New natural products of peptidepolyketide glycoside totopotensamide A and its aglycone totopotensamide B were isolated from the mollusk-associated marine actinobacteria of Streptomyces sp. (Lin et al., 2012). In addition, a new potential thiazolyl peptide called kocurin was isolated from Kocuria sp. and Microccocus sp. (sponge-derived) and showed antimicrobial potential against human bacterial pathogens (Palomo et al., 2013). Ascidian (Aplidium lenticulum)–associated marine Streptomyces sp. JP90 produced a new organophosphate cinnamoylphosphoramide that comprises a rare natural phosphoramide methyl ester, which inadequately inhibited acetylcholinesterase (Quitschau, Schuhmann, Piel, Von Zezschwitz, & Grond, 2008). Sponge-associated

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Actinomadura sp. produced the Diterpene JBIR-66 compound, and it showed uncertain radical scavenging activities (Takagi et al., 2010). Streptomyces sp. isolated from demospongiae sponge produced the salicylamide derivative compound, which showed moderate toxicity against HeLa cells (Ueda et al., 2009). Unidentified tunicate-associated marine actinobacteria Streptomyces sp. BOSC-022A produced the Barmumycin compound, and it showed cytotoxic activity against various human tumor cell lines (Lorente, Pla, Canedo, Albericio, & Alvarez, 2010). In addition, a new tetromycin bioactive compound was isolated from the Mediterranean sponge–associated actinobacteria Streptomyces axinellae Pol001 (Pimentel, Scheuermayer, Kozytska, & Hentschel, 2009). These tetromycins showed activity against the human breast cancer cell line MDA-MB 435 and Try human liver cancer cell line 7402, respectively (Wei et al., 2011). The new teleocidin analog, the JBIR-31 compound, was isolated from Haliclona sp. marine sponge–associated Streptomyces sp. NBRC 105896, and it showed a weak cytotoxic effect against HeLa cells (human cervical carcinoma) with IC50 value of 49 mM (Hosoya, Hirokawa, Takagi, & Shinya, 2012).

5. CONCLUSION Marine actinobacteria are the resource for the production of biological active compounds. Around 70% of marine natural products are isolated from marine bacteria alone and marine organism–associated actinobacterial enzymes and other metabolites. It could provide different biological activities, such as antimicrobial activity, cytotoxicity, antioxidant activity, antiHIV activity, and anticancer activities. Moreover, pharmacological potential enzymes from terrestrial microorganisms have not been extensively reported. However, widely scattered marine microorganisms are making a strong case for future investigation of marine bacteria for biodiscovery purposes.

ACKNOWLEDGMENTS The corresponding author (Kamala) would like to acknowledge and thank the Science and Engineering Research Board-Department of Science and Technology, Government of India, for providing fund through the National Post-Doctoral Fellowship program (File No: PDF/2015/000680). The authors are also grateful to the Center for Environmental Nuclear Research, Directorate of Research, SRM University, Tamil Nadu, for providing facilities.

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CHAPTER SIX

Production of Enzymes From Agricultural Wastes and Their Potential Industrial Applications S. Bharathiraja*, J. Suriya†, M. Krishnan†, P. Manivasagan{, S.-K. Kim{,§,1 *CAS in Marine Biology, Annamalai University, Porto Novo, India † School of Environmental Sciences, Bharathidasan University, Tiruchirappalli, 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 address: [email protected]; [email protected]

Contents 1. Introduction 2. Enzymes Production From Agricultural Wastes 2.1 Amylase 2.2 Cellulase 2.3 Tannase 2.4 Xylanase 2.5 Protease 2.6 Laccase 3. Actinobacterial Enzymes Produced From Agricultural Wastes 4. Conclusion Acknowledgments References

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Abstract Enzymatic hydrolysis is the significant technique for the conversion of agricultural wastes into valuable products. Agroindustrial wastes such as rice bran, wheat bran, wheat straw, sugarcane bagasse, and corncob are cheapest and plentifully available natural carbon sources for the production of industrially important enzymes. Innumerable enzymes that have numerous applications in industrial processes for food, drug, textile, and dye use have been produced from different types of microorganisms from agricultural wastes. Utilization of agricultural wastes offers great potential for reducing the production cost and increasing the use of enzymes for industrial purposes. This chapter focuses on economic production of actinobacterial enzymes from agricultural wastes to make a better alternative for utilization of biomass generated in million tons as waste annually.

Advances in Food and Nutrition Research, Volume 80 ISSN 1043-4526 http://dx.doi.org/10.1016/bs.afnr.2016.11.003

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1. INTRODUCTION Globally five billion metric tons of biomass has been produced annually from agriculture including ground nut cake, rice bran, rice straw, sugarcane bagasse, fruits and vegetable wastes, wheat bran, cotton leaf scraps, etc. (http://www.unep.org/gpwm/FocalAreas/WasteAgriculturalBiomass/tabid/ 56456/Default.aspx). Henceforth, it is very crucial to convert these wastes effectively and economically into valuable products of industrial and commercial potential and also to reduce the detrimental impact of these wastes on environment (Wang et al., 2016). Lignocellulose and starch are the major content of the wood industry, agroindustry, and domestic and garden wastes. About half of the plant matter is composed of lignocelluloses which is the most abundant renewable organic matter in soil (Singh, Kapoor, & Kumar, 2012). It composed of cellulose (35–50%), hemicellulose (20–35%), and lignin (15–25%) that is strongly connected together by variety of noncovalent and covalent linkages (Limayem & Ricke, 2012). Cellulose is the most common carbohydrate in plants that consists of linear biopolymer of an hydroglucose units linked by the β-1,4-glycosidic bond. It is very difficult to degrade these components rapidly in the natural environment (Mondragon et al., 2014). Endoglucanase, exoglucanase, cellobiohydrolase, and β-glucosidase act together to hydrolyze cellulose (Tao, Zhu, & Huang, 2010). Hemicelluloses are heteropolysaccharides consist of D-glucose, D-galactose, D-xylose, L-arabinose, or D-mannose sugars and may contain xylans, glucans, mannans, arabinoxylans glucuronoxylans, β-glucans, glucomannans, galactomannans, galactoglucomannans, and xyloglucans (Limayem & Ricke, 2012; Scheller & Ulvskov, 2010). The composition and branching vary among different plant sources. Xylan is the most abundant hemicellulosic component composed of the polymer of xylose (Limayem & Ricke, 2012). Hemicellulose is degraded by the synergetic action of different xylanolytic enzymes such as endo-1,4-β-xylanase and xylan 1,4-β-xylosidase (Rozanov et al., 2015). Xylanase hydrolyzes β-1,4 linkages of the xylan molecule, whereas β-xylosidase cleaves xylobiose and xylooligosaccharides into xylose. Lignin is a complex heteropolymer of polyphenols composed of sinapyl alcohol pcoumaryl alcohol and coniferyl alcohol (Agbor, Cicek, Sparling, Berlin, & Levin, 2011). Lignin gives structural support to the plants and also acts as an impermeable barrier to the enzymes (Brodeur et al., 2011). Starch is a polysaccharide where 20–25% of the molecule is composed of amylose consisting of linear chain of glucose units joined by α-1,4-glycosidic

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linkage. Near 75–80% of starch is occupied by amylopectin which consist of branched chains of glucose units. α-1,4-Glycosidic linkage connects the linear glucose units, whereas branched glucose units are united together by α-1,6-glycosidic bonds. Temperature, hydrolysis conditions, and enzyme origin have the major impact on the end product of starch hydrolysis (http://rcsb.org/pdb/images/1hny_bio_r_500.jpg?bioNum¼1). Amylases are enzymes able to hydrolyze starch producing dextrins and glucose (Homaei, Ghanbarzadeh, & Monsef, 2016). Cellulases are utilized for biofuel, textile, pulp, paper, detergent industries, animal feed, and food (Sukumaran, Singhania, & Pandey, 2005) as well as in ligand-binding studies (Gupta, Samant, & Sahu, 2012). Hemicellulases are applied in clarification of fruit juices, paper waste deinking, biobleaching, and to improve the quality of feed, fodder, and fibers as well as for saccharification of hemicelluloses to xylose sugars (Soni & Kango, 2013). Lignin-degrading enzymes are widely used for the pretreatment of recalcitrant lignocellulosic biomass for biofuel production and also applied in textile, food, paper, cosmetic, pharmaceutical industries, organic synthesis, wastewater treatment, and bioremediation (Abdel-Hamid, Solbiati, & Cann, 2013). Amylases have been utilized in textile, brewing, detergent, pharmaceuticals, sugar, paper, and distilling industries (de Souza & Magalha˜es, 2010). The utilization of biological wastes for growing microorganisms may constitute a better alternative for enzyme production with lower costs. Large number of commercial enzyme preparations from bacteria and fungi has limitations in their activity to degrade biomass with respect to several parameters including pH, temperature, etc. Actinobacteria are distinctive prokaryotic organisms which have the characteristic features of both bacteria and fungi. It has immense potential for producing bioactive molecules and is promising source of industrially essential enzymes, owing to their greater metabolic diversity and higher adaptability to extreme environmental conditions (Chakraborty et al., 2014).

2. ENZYMES PRODUCTION FROM AGRICULTURAL WASTES Microbial enzymes are more stable and active compared to plant and animal enzymes; henceforth, they have wide range of medicinal and industrial applications. Moreover, their biochemical diversity, sensitivity to gene manipulation, and large-scale production in a short duration by

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fermentation make microorganisms as an alternative source for the enzyme production. Therefore, industries are in need of novel microbial strains for various enzyme productions to meet the current enzyme requirements. In this chapter, we will highlight the application of lignocellulosic wastes as substrates for microbial growth and to produce industrially valuable enzymes such as amylase, cellulases, tannase, xylanase, protease, and laccase by microorganisms except actinobacteria that will be discussed later.

2.1 Amylase Many agricultural wastes such as rice bran, rice straw, rice straw + rice bran, wheat bran, red gram husk, jowar straw, and jowar spathe have been tested for the production of α-amylase from Gibberella fujikuroi. Maximum production of α-amylase was observed in wheat bran (Mulimani & Ramalingam, 2000). Wheat bran and rice husk were tested for α-amylase production by Bacillus subtilis (Baysal, Uyar, & Aytekin, 2003), and 7.3-fold higher enzyme production in wheat bran compared to rice husk was reported. Francis, Sabu, Nampoothiri, Ramachandran, et al. (2003) used spent brewing grains as the sole carbon source for α-amylase secretion by Aspergillus oryzae and found 20% increase in enzyme yield under optimized conditions. Soni, Kaur, and Gupta (2003) obtained very high titers of thermostable— α-amylase by Bacillus sp. using wheat bran. Ramachandran, Patel, Nampoothiri, Francis, et al. (2004) produced α-amylase from A. oryzae using coconut oil cake as a substrate. High titers of α-amylase was produced by Bacillus sp. in wheat bran with the addition of nutrients like soya bean meal, glycerol, and MgSO4  7H2O (Sodhi, Sharma, Gupta, & Soni, 2005). Two α-amylases, AmyI (76 kDa) and AmyII (53 kDa), were produced by the thermophilic Bacillus sp. strain WN11. The percent hydrolysis of raw potato and corn starch was 77% and 44% for AmyI and 82% and 37% for AmyII (Mamo & Gessesse, 1999). Vijayaraghavan, Kalaiyarasi, and Vincent (2015) used cow dung as the potential carbon source for the production of amylase from Bacillus cereus. They obtained maximal amylase production in the presence of 0.01% ammonium sulfate, 0.1% fructose, and 100% moisture. Mihajlovski, Radovanovic, Veljovic, Siler-Marinkovic, and Dimitrijevic-Brankovic (2016) used molasses and sugar beet pulp for improved β-amylase production by Paenibacillus chitinolyticus CKS1. Under optimized conditions (3% sugar beet pulp concentration 10% inoculum concentration and 88.07 h of incubation time 83.07 h), they obtained highest production. Ikram-Ul-Haq, Mahmood, and Javed (2012) used

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agroindustrial wastes for the production of α-amylases from Paenibacillus amylolyticus. Jin, van Leeuwen, Patel, and Yu (1998) utilized starch processing wastewater for α-amylase production from A. oryzae. Orange waste powder was used as sole carbon source for α-amylase production by Aspergillus niger ATCC 16404 (Djekrif-Dakhmouche, Gheribi-Aoulmi, Meraihi, & Bennamoun, 2006). Potato starchy waste, wheat bran, and rice husk were used as a costeffective carbon substrate for amylase enzyme production by B. subtilis (Asgher, Asad, Rahman, & Legge, 2007; Baysal et al., 2003; Shukla & Kar, 2006). Abd-Elhalem, El-Sawy, Gamal, and Abou-Taleb (2015) employed potato starchy waste as the sole carbon source for enhanced production of amylase enzymes and also found that α-, β-, γ-amylases activity was increased by Bacillus amyloliquefaciens about 1.26-, 4-, and 8-fold, respectively, after 48 h at 50°C using rotary shaker. Singh and Gupta (2014) produced and characterized amylase enzyme from Aspergillus flavus TF-8 using Sal (Shorea robusta) deoiled cake.

2.2 Cellulase Cellulolytic microorganisms are mostly utilizing carbohydrates for their energy but are unable to use proteins or lipids as energy source for their growth (Lynd, Weimer, Van Zyl, & Pretorius, 2002). Picart, Diaz, and Pastor (2007) produced high level of cellulase in media supplemented with rice straw by Penicillium sp., and they found that the enzyme was optimally active and stable at 65°C and pH 4–5. Annamalai, Rajeswari, and Balasubramanian (2014) reported higher activities of cellulase enzyme (4040.45 U/mL) with 6.27 g/L rice bran, 2.52 g/L of yeast extract, an initial pH 9.0, and temperature 50°C in Bacillus carboniphilus CAS 3. Trichoderma reesei and its mutant were cocultured with Aspergillus phoenicis QM329 for cellulase production on bagasse. Parent Trichoderma strain and the Aspergillus revealed the synergistic action, resulting in enhanced production of cellulase, endoglucanase, and β-glucosidase activities (Gutierrez-Correa & Tengerdy, 1997). Liu et al. (2011) grown Aspergillus fumigates Z5 under solid-state fermentation (SSF) for thermostable cellulase production to degrade agricultural wastes for ethanol production. It yields 0.112 g bioethanol/g dry substrate, suggesting that it is a promising fungus in the bioethanol production process. Gautam et al. (2011) used municipal solid waste (MSW) residue for the production of cellulase by A. niger and Trichoderma sp. The results revealed

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that MSW residue (4.0%) was the best carbon substrate for endoglucanase (1.95 U/mL), exoglucanase (1.77 U/mL), and β-glucosidase (1.66 U/mL) by Trichoderma sp. than A. niger. The maximum production of carboxymethyl cellulase (102 U/mL) from B. amyloliquefaciens DL-3 with 2% rice hull and 0.25% peptone as the carbon and nitrogen sources and they also found that rice hull and rice bran were better carbon sources for the enzyme production than glucose, fructose, maltose, and sucrose (Jo, Lee, Kim, et al., 2008). Waghmare, Kshirsagar, Saratale, Govindwar, and Saratale (2014) investigated the potential of various agricultural wastes such as sugarcane bagasse, sugarcane barbojo, sorghum husks, grass powder, corn straw, and paddy straw for cellulolytic enzyme production by Klebsiella sp. PRW-1. Grass powder and sugarcane barbojo were found to be the best carbon sources for enzyme production. dos Santos, Gomes, Bonomo, and Franco (2012) used potato peel for the production of cellulolytic enzymes from A. niger. Pleurotus ostreatus and Pleurotus sajor-caju were evaluated for their ability to produce various lignolytic and cellulolytic enzymes on banana agricultural wastes (Reddy, Ravindra Babu, Komaraiah, Roy, & Kothari, 2003). For cellulase production by Trichoderma harzianum, temperature, pH, inoculums concentration, agitation rate, substrate (domestic wastewater sludge), and cosubstrate (wheat flour) were optimized using two-level fractional factorial design. They obtained highest cellulase activity (10.2 Filter Paper Unit (FPU)/mL) during the fermentation process which was 1.5-fold higher than the cellulase produced from the results of design of experiment (6.9 FPU/mL) (Alam, Muyibi, & Wahid, 2008). Li, Liu, Bai, and Zhao (2016) used fed-batch strategy for overproduction of cellulase from T. reesei RUT C3, through which glucose was controlled at low level. Cellulase activity as high as 90.3 FPU/mL was obtained at 144 h, and cellulase productivity was increased to 627.1 FPU/L/h. Krishna (1999) utilized banana wastes for cellulase production by B. subtilis (CBTK 106). An extracellular halotolerant, thermoalkaline cellulase produced by Bacillus halodurans CAS 1 from lignocellulosic wastes with an optimum pH, temperature of 9.0 and 60°C, respectively. The enzyme activity was greatly inhibited by the addition of EDTA and PMSF revealed that it was a metalloenzyme and serine residue was essential for its catalytic activity (Annamalai, Rajeswari, Elayaraja, & Balasubramanian, 2013). Aspergillus spp. MPS-002 and Phyllosticta spp. MPS-001 were evaluated for their ability for producing lignocellulolytic enzymes on banana agricultural waste (Shah, Reddy, Banerjee, Babu, & Kothari, 2005). Sporothrix carnis was grown on corncob to produce thermostable crude cellulase. Maximum production of cellulase

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(285.7 U/mL) was obtained at 96 h of cultivation with 2.5% inoculum at pH 6.0, temperature 80°C (Olajuyigbe & Ogunyewo, 2016).

2.3 Tannase Glucose and gallic acid are produced from tannic acid by the hydrolytic potential of tannase. Tannases are mostly applied in the food and pharmaceutical industries. Tannase is also utilized in the stabilization of malt polyphenols, treatment of tannery effluents, clarification of beer and fruit juices, and it also prevents the madeirization in fruit juices and wine (Aguilar & Sanchez, 2001). Sabu, Pandey, Daud, and Szakacs (2005) utilized palm kernel cake and tamarind seed powder for the production of tannase by A. niger. Palm kernel cake produced 13.03 IU/g ds and tamarind seed powder yield 6.44 IU/g ds. Kumar, Sharma, and Singh (2007) produced tannase from Aspergillus ruber using different tannin-rich substrates like amla leaves (Phyllanthus emblica), jamun leaves (Syzygium cumini), ber leaves (Ziziphus mauritiana), and jawar leaves (Sorghum vulgaris). They observed that jamun leaves produced high titers of tannase (69 U/g ds) than others. Banerjee, Mondal, and Pati (2007) used various agricultural by-products such as rice bran, wheat bran, rice straw dust, saw dust, and sugarcane pith as sole substrates for tannase production by Aspergillus aculeatus. Maximum production of tannase was observed in wheat bran. Nandini, Nandini, and Krishna Sundari (2014) used food and agriculture residue for maximum tannase production. They found that highest tannase (13.21 U/g) and gallic acid (3.51 mg/g) production by B 2.2 bacterial isolate followed by 9.15 U/g of tannase and 3.36 mg/g of gallic acid by B 2.7 bacterial isolate. de Lima et al. (2014) agroindustrial waste for tannase production from Penicillium montanense URM6286 under SSF. It revealed maximum tannase activity (41.64 U/mL) after 72 h of incubation with barbados cherry at pH 9.0 and 50°C. It also stable over a wide pH range and temperature. It was applied for grape juice clarification, showed higher efficiency by reducing 46% of the tannin content after 120 m of incubation. The fungal strain Penicillium atramentosum KM produced extracellular tannase under submerged fermentation (SmF) using amla jamun, ber, keekar, and jamoa powdered leaves. Among these substrates, amla (2%, w/v) and keekar leaves (3%, w/v) revealed highest tannase production (32.8 and 34.7 U/mL, respectively) (Selwal & Selwal, 2012). Different agroindustrial wastes such as red gram husk, coffee husk, coconut powder, green gram husk, ground nut waste, cotton seed waste, wheat

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bran, tamarindus seed powder, rice bran, Cicer arietinum, cashew apple bagasse, and corn powder were used for tannase production by Aspergillus terreus. Among the wastes, wheat bran supported maximum tannase production with an optimum pH of 3.5 after 72 h of incubation (Malgireddy & Nimma, 2015). Klebsiella pneumonia KP715242 was used by Kumar, Singh, Beniwal, and Salar (2016) to obtain maximum tannase production from agroresidues like Indian gooseberry leaves, black plum leaves, Eucalyptus leaves, Eucalyptus globulus, and Babul leaves. Among all agroresidues, Indian gooseberry leaves were found to be the best substrate for the production of tannase under SmF. Bhoite and Murthy (2015) used coffee pulp as substrate for maximum also production of tannase by Penicillium verrucosum at 96 h of fermentation period. Mohan, Viruthagiri, and Arunkumar (2014a, 2014b) applied RSM for the highest production of tannase by Aspergillus foetidus (MTCC 3557) using red gram husk as substrate under SmF with an optimum pH of 5.5, temperature of 35.5°C, tannin content of 3.1%, and fermentation period of 97 h. Varadharajan, Vadivel, Ramaswamy, Sundharamurthy, and Chandrasekar (2015) investigated various agrowastes as substrates for the production of tannase production from A. oryzae under SmF and found pomegranate rind extract as the best substrate with a tannase yield of 138.12 IU/mL. Several natural substrates, such as aggase, ground nut oil cake, rice bran, tamarind seed powder, wheat bran (Natarajan & Rajendran, 2012), tea residue, and coffee pulp (Bhoite & Murthy, 2015; Sharma et al., 2014), have been used as a sole carbon source for maximum tannase production under SSF.

2.4 Xylanase Xylanases are widely utilized for clarification of juice and wine, biobleaching of kraft pulp, improving the bakery product’s quality, and in animal feed (Bhat, 2000). Gawande and Kamat (1999) evaluated many lignocellulosic residues such as wheat bran, sugarcane bagasse, rice straw, and soya bean hulls for xylanase production from A. terreus and A. niger. Highest xylanase production was observed in wheat bran. This was a remarkable result because wheat bran is cheaper than xylan used for xylanase production and furthermore, both Aspergillus strains produced xylanase with lower levels of cellulase making the purification processes much easier. Milagres, Santos, Piovan, and Roberto (2004) showed the production of high level of thermostable xylanase (1597 U/g xylanase activity) after 10 days of SSF from Thermoascus aurantiacus. Sonia, Chadha, and Saini (2005)

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produced maximum levels of cellulase-free xylanase (11,855 U/g dry substrate) and associated hemicellulases by thermophilic isolate Thermomyces lanuginosus grown on sorghum straw. Virupakshi, Gireesh Babu, Gaikwad, and Naik (2005) tested various easily available lignocellulosic substrates for xylanase production by Bacillus sp. and found that rice bran was the best substrate for xylanase production. Botella, Diaz, de Ory, Webb, and Blandino (2007) revealed the immense potential of grape pomace to produce xylanase by Aspergillus awamori. Mamma, Kourtoglou, and Christakopoulos (2007) used orange peels to produce high xylanase activities from A. niger. Patel and Prajapati (2014) investigated different agricultural wastes such as wheat husk, rice bran, and rice straw for xylanase production by using Cladosporium sp., and he found that maximum xylanase activity in rice bran. Ho (2015) used solidstate and submerged fermentation techniques for xylanase production from B. subtilis ATCC 6633 using inexpensive agricultural wastes. Higher xylanase activity of 11.646  4.163 U/mL was obtained with barley husk in SmF. Whereas wheat bran in SSF produced 22.071  0.186 U/mL of xylanase activity at 48 h of fermentation. Azin, Moravej, and Zareh (2007) also reported combination of wheat bran and wheat straw at the ratio of 7:3 produced the maximum xylanase activity of 479.7 U/g by Trichoderma longibrachiatum. Gandarillas, Soto, and Vargas (2012) used barley straw by Bacillus spp. Lb-4 and they found that maximum xylanase activity of 10 U/mL. Haddar, Driss, Frikha, Ellouz-Chaabouni, and Nasri (2012) also reported that the maximum xylanase activity of 18.66 g/L was produced by Bacillus mojavensis A21 using barley bran. Soliman, Sherief, and El-Tanash (2012) investigated the production of xylanase from A. niger and Trichoderma viride using several agricultural residues. They reported that barely bran was the suitable substrate for highest enzyme production (42.5 U/g substrate) from T. viride after 2 days of incubation, at pH 5.5 and temperature 35°C. Herculano et al. (2016) reported the maximum xylanase activity (29,085  1808 U g/g dry substrate) with 5.0 g of substrate, initial moisture of 15% at 25°C and pH 6.0, after 120 h of fermentation from Aspergillus japonicus URM5620 using SSF of castor press cake (Ricinus communis). Various agroindustrial wastes such as passion fruit peel brewer’s spent grain, soybean waste, wheat bran, soybean peel, corn straw, orange peel, pineapple peel, apple peel, sugarcane straw, and sugarcane bagasse were investigated for the production of xylanase enzyme by an A. niger. Maximum xylanase activity was obtained at pH 5.0, at 30°C, during 5 days of incubation in brewer’s spent grain (Izidoro & Knob, 2014).

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Ho and Chinonso (2016) used ethyl methane sulfonate and acridine orange to produce random mutant of B. subtilis to increase the xylanase production using barley husk as the substrate by SmF. Kanimozhi and Nagalakshmi (2014) obtained highest xylanase production of 1.4 U/mL in wheat bran as substrate.

2.5 Protease Proteases are used in several industries such as pharmaceutical, food, detergent, silk, and leather and also for recovery of silver from used X-ray films (Jisha et al., 2013). Germano, Pandey, Osaku, Rocha, and Soccol (2003) utilized defatted soya bean cake as a support-substrate for the production of protease by Penicillium sp. The crude enzyme retains their activity (50–60%) in the presence of commercial detergents. Tunga, Shrivastava, and Banerjee (2003) produced an extracellular alkaline serine protease from Aspergillus parasiticus using wheat bran as a support-substrate. The produced protease was stable and active in various detergents. Uyar and Baysal (2004) evaluated the efficiency of wheat bran and lentil husk for the production of alkaline protease by Bacillus sp. Obtaining maximum enzyme levels of 429.041 and 168.640 U/g, respectively. Sandhya, Sumantha, Szakacs, and Pandey (2005) compared the potential of various agroindustrial wastes like coconut oil cake, rice bran, spent brewing grain wheat bran, rice husk, palm kernel cake, sesame oil cake, olive oil cake, and jackfruit seed powder to produce neutral protease by A. oryzae. They found that wheat bran was the best producer of protease. Prakasham, Rao, and Sarma (2006) studied the alkaline protease production by Bacillus sp. using different agroindustrial waste materials (wheat bran, red gram, green gram, black gram, chick pea, and husks). They observed the highest production of enzyme in green gram husk. Mahanta, Gupta, and Khare (2008) found that deoiled Jatropha seed cake was a best source for protease (1818 U/g substrate) and lipase production (625 U/g). de Castro, Soares, Albernaz, and Sato (2016) produced surfactant, solvent, salt, and oxidizing agent tolerant protease from A. niger using different agricultural wastes. Shivasharanappa, Hanchinalmath, Sai Sundeep, Borah, and Prasad Talluri (2014) isolated Trichoderma viridiae strain VPG 12 from agriculture soil and produced alkaline protease from this novel strain using various agricultural wastes such as red gram husk, green gram husk, and Bengal gram husk. Protease production and its optimization in surface-modified coffee pulp waste and corncobs using Bacillus sp. by SSF were reported by Kandasamy et al. (2016). They obtained the maximum

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activity of protease (920 U/mL) at 60 h of incubation with 3.0 g/L of coffee pulp waste and 2.0 g/L of corncobs with an optimum pH 8 and temperature 37°C. Sankareswaran, Anbalagan, and Prabhavathi (2014) studied the production and optimization of protease produced by Bacillus using agricultural wastes in both solid-state and SmF. de Castro, Nishide, and Sato (2014) used kinetic and thermodynamic parameters for maximum production of protease by A. niger LBA02 under SSF using different agroindustrial by-products as sole carbon source and also reported its biochemical characterization. Sathishkumar, Ananthan, and Arun (2015) isolated protease producing B. subtilis GACAS8 from marine ascidian Phallusia arabica and obtained the maximum protease activity at 50°C and pH 9. Molecular weight of the purified protease was 41 kDa, and its activity was stimulated by Mg2+ and Ca2+, strongly inhibited in the presence of EDTA, and it revealed resistance to Tween 20, Tween 40, and SDS. de Castro and Sato (2014) reported about production and characterization of protease from A. oryzae and also they evaluated the physical–chemical parameters using different agro-industrial wastes. de Castro and Sato (2013) investigated the combined effects of agroindustrial wastes on the production of protease and α-amylase at the same time under SSF, and they found that highest protease and α-amylase activities were obtained wheat bran as the substrate. Bajaj, Sharma, and Singh (2013) reported the enhanced production of fibrinolytic protease from B. cereus NS-2 using cotton seed cake as nitrogen source. Murthya and Kusumoto (2015) used potato pulp powder for acid protease production from A. oryzae. Meena, Tripathi, Srivastava, and Jha (2013) utilized wheat bran for alkaline protease production using Pseudomonas aeruginosa under SSF. Abraham, Gea, and Sa´nchez (2013) evaluated the potential of soy fiber residues for alkaline protease production. The highest activity was of 47,331  1391 U/g dry matter for soy fiber, and 18,750  1596 U/g dry matter for soy fiber with 10% compost was attained under thermophilic conditions. Thanapimmetha, Luadsongkrama, Titapiwatanakun, and Srinophakun (2012) used Jatropha curcas residue for protease production by A. oryzae.

2.6 Laccase Laccases have been widely applied in various industries owing to their following features: broad substrate specificity; most enzymes are extracellular;

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henceforth, it is very easy to purify the enzyme, it does not require the addition of cofactor, and it generally exhibits a considerable level of stability and adaptability in the extracellular environment. Such characteristics make laccases most preferred enzyme for several industrial applications such as biopulping, biobleaching, and the treatment of industrial wastewater. Most of the agricultural wastes are rich in soluble carbohydrates and also have laccase synthesis inducers which resulting in high production of laccase (Rosales, Rodrı´guez Couto, & Sanroma´n, 2005). Rodrı´guez Couto and Sanroman (2005) reported the maximum laccase activities using agrowastes as support-substrate, in particular those rich in cellulose. Osma, Toca Herrera, and Rodrı´guez Couto (2007) have reported the alternative substrates for efficient production of laccase at low cost. Rebhun, Wasser, and Hadar (2005) utilized agroindustrial waste for laccase production by white-rot basidiomycetes such as Cerrena unicolor and Cucurbita maxima, and they found that wheat bran was the best growth substrate for fermentation of C. unicolor, enabling a very high production of laccase (87.450 IU/L on day 7). do Rosario Freixo, Karmali, Fraza˜o, and Arteiro (2008) used Coriolus versicolor for laccase production using tomato pomace as sole carbon and found that the highest laccase activity (362 U/L of fermentation broth) after third day of incubation. Avanthi and Banerjeea (2016) studied the laccase-mediated lignin degradation of lignocellulosic feedstocks for ethanol production. Lu et al. (2013) characterized the laccase-like multicopper oxidase from Streptomyces sp. C1 in agricultural waste compost and applied it for decolorization of azo dyes. Wang, Hu, Guo, and Liu (2014) reported the increased laccase production by Trametes versicolor using corn steep liquor as both nitrogen source and inducer. Gonzalez et al. (2013) used submerged and semisolid culture conditions for laccase production from Trametes pubescens on agroindustrial wastes. Karp et al. (2012) characterized laccase isoforms of P. ostreatus in SSF using sugarcane bagasse. Birhanli and Yes¸ilada (2013) utilized lignocellulosic wastes such as sunflower receptacle, apricot seed shell, and bulrush for laccase production under semisolid-state and SmF conditions from Trametes trogii (Berk.) ATCC 200800 and T. versicolor (L.) ATCC 200801. Risdianto, Sofianti, Suhardi, and Setiadi (2012) investigated the optimum conditions for laccase production from white-rot fungi such as Marasmius sp., Trametes hirsuta, T. versicolor, and Phanerochaete chrysosporium on empty fruit bunches, rice straw, corncob, and rice husk. Periasamy, Rajadurai, and Palvannan (2011) analyzed the impact of various agrowastes on laccase production

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by P. ostreatus IMI 395544. Karp et al. (2015) optimized the production of laccases and evaluated the delignification of sugarcane bagasse by P. ostreatus in SSF.

3. ACTINOBACTERIAL ENZYMES PRODUCED FROM AGRICULTURAL WASTES Most agricultural wastes generated in field and processing sites are discarded as waste, creating environmental problems, if not properly disposed. The main aim of biotechnology is to produce industrially pivotal enzymes at low cost. Among microbial community, actinobacteria are considered as an efficient group that degrades plant biomass in nature and further most of the actinobacteria have important features like thermostability and adaptability to extreme environmental conditions. Henceforth, large number of industrially pivotal enzymes with distinct properties can be produced. Therefore, we have separated this section to reveal the crucial role of actinobacteria in lignocellulolytic enzyme production. Streptomyces sp. Ab106 was tested for the production of cellulase-free xylanase on sugarcane bagasse. Central composite experimental design was used to evaluate the impact of pH and temperature on the xylanase production. The highest yield of xylanase without cellulase and mannanase activities was obtained at 50°C and pH 7.0 (Techapun, Charoenrat, Watanabe, Sasaki, & Poosaran, 2002). de Lima, do Nascimento, da Silva Bon, and Coelho (2005) produced cellulase by Streptomyces drozdowiczii using agroindustrial by-products. The enzyme showed resistance to commercials detergents. Moreover, the potential utility of this enzyme in textile processing was tested by comparing with commercial enzyme (IndiAge Super L). Ponce-Noyola and de la Torre (2001) compared the filter paperase and xylanase production by wild and catabolic depressed mutant strains of Cellulomonas flavigena on sugarcane bagasse. Catabolic depressed mutants produced remarkably higher levels of enzymes than wild strain. Sugarcane bagasse induced both wild and mutant strains to produce three to eight higher levels of filter paperase and xylanase. Cellulase-free xylanase was produced by Streptomyces sp. Abl06 using cane bagasse. It produced maximum enzyme activity without cellulase and mannanase activities at pH 6.0 and 60°C, and it retained its 70% activity at 60°C at pH 9 (Techapun, Charoenrat, Poosaran, Watanabe, & Sasaki, 2002). The production optimization of α-amylase by Streptomyces sp. 20r using an agroindustrial residue (orange waste) as the primary carbon source was

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performed on Plackett–Burman experimental designs. Statistical analysis showed that the changes in substrate concentration (5–15%), NaCl (0–6.5%), inoculums concentration (5–10%), and pH (5–9) produced highest α-amylase (8.26 U/mL) in SmF (Mounaimen & Mahmoud, 2015). Padmavathi, Thiyagarajan, Naveed Ahamed, and Palvannan (2011) evaluated the production of xylanase by Streptomyces coelicolor using four different substrates such as sugarcane bagasse, pineapple, orange peels, and pomegranate peels and found that the sugarcane bagasse was the best substrate for xylanase production. Rice straw pulp was used as a low-cost substrate for xylanase production by Streptomyces albus and Streptomyces chromofuscus. They obtained maximum xylanase activity on pulp treated with TiO2 and untreated rice straw pulp in both Streptomyces species, respectively (Rifaat, Nagieb, & Ahmed, 2005). Brito-Cunha, Gama, Jesuino, Faria, and Bataus (2015) analyzed the cellulases from Streptomyces thermocerradoensis I3 using sugarcane bagasse or wheat bran. The highest carboxylmethyl cellulase activity was obtained when the microorganisms were cultivated on wheat bran supplemented medium. The effect of the carbon source (sisal waste and sugarcane bagasse) on α-amylase production by Streptomyces sp. SLBA-08 was studied using submerged cultivations at 30°C. The highest level of α-amylase activity corresponded to 10.1 U/mL and was obtained using sisal waste (2.7%) (dos Santos, Teles, et al., 2012). Agroindustrial wastes such as wheat straw, wheat bran, sugarcane bagasse, rice bran, and corncob were used for the production of xylanase and amylase by Streptomyces sp. The amylase and xylanase activity were maximum in rice bran (Singh et al., 2012). Streptomyces sp. produces xylanase on various feed stuffs like oat spelt xylan, wheat bran, sugarcane molasses, tomato pomace, rice bran, and saw dust under SmF condition. Out of all lignocellulosic waste, the highest amount of xylanase enzyme was produced in oat spelt xylan medium (Kalpana & Rajeswari, 2015). Bhosale, Sukalkar, Uzma, and Kadam (2011) studied the potential of cheap agricultural sources such as corncobs, wheat bran, sugarcane bagasse, and rice bran as substrate in liquid shake culture medium for xylanase production by Streptomyces rameus. Maximum level of xylanase activity was observed on 20 g/L sugarcane bagasse at 30–50°C and pH 8.5. Rathnan and Ambili (2011) used Streptomyces sp. for cellulase production from fruit waste. They obtained highest enzyme production at pH 5.0 and 40°C. Dilipkumar, Rajasimman, and Rajamohan (2013) used coconut oil cake as a sole carbon source to produce inulinase by Streptomyces sp. Manivasagan, Venkatesan, Sivakumar, and Kim (2013) produced and

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characterized protease having antioxidant efficiency from Streptomyces sp. MAB18 using poultry wastes. Specific activity of this protease was 2398.36 U/mg and the molecular weight was estimated as 43 kDa. Streptomyces misionensis PESB-25 secreted thermoacidophilic endoglucanase from sugarcane bagasse and corn steep liquor added medium. A peak of endoglucanase accumulation was observed in a medium with sugarcane bagasse 1.0% (w/v) and corn steep liquor 1.2% (w/v) within 3 days of cultivation (Franco-Cirigliano et al., 2013). Macedo et al. (2013) evaluated the potential of sisal bagasse, sugarcane bagasse, and straw waste for cellulase enzyme from Streptomyces sp. The results showed that maximum carboxymethyl cellulase production (1.11 U/mL) was obtained on 2.4% (w/v) sisal bagasse and 0.3% (w/v) ammonium sulfate as the carbon and nitrogen sources after 48 h of incubation. Napier grass, sugarcane bagasse, and soybean bran were screened for xylanase production from Streptomyces viridosporus T7A. The maximum xylanase production (423.9 U/g) was obtained in the medium supplemented with 10% soybean meal, 20% Napier grass, and 70% sugarcane bagasse (Alberton et al., 2009). Sugarcane bagasse or wheat bran was used as carbon source, and corn steep liquor used as nitrogen source for cellulase production from Streptomyces viridobrunneus SCPE-09. The results revealed that 2.0% wheat bran (w/v) and 0.19% corn steep liquor (w/v) gave highest production of carboxymethyl cellulase, after 5 days of incubation (Da Vinha et al., 2011). Nascimento, Junior, Pereira, Bon, and Coelho (2009) obtained maximum cellulase production by Streptomyces malaysiensis occurred within 4 days incubation when using a growth medium containing brewer’s spent grain 0.5% (w/v) and corn steep liquor 1.2% (w/v). Carboxymethyl cellulases activity showed to be stable over an acid pH range (2.0–7.0) and in temperatures of 40–60°C. Kar, Ray, and Mohapatra (2008) used various agricultural wastes such as cassava bagasse, sugarcane bagasse, and wheat bran for the amylase production from Streptomyces erumpens MTCC 7311, and they reported that wheat bran was found to the best substrate. Untreated ballmilled bagasse was used for cellulase production from three actinomycetes such as Streptomyces albogriseolus, Streptomyces nitrosporeus, and Micromonospora melanosporea. These mesophilic actinomycetes simultaneously secreted cellulases, xylanases, β-xylosidases (Van Zyl, 1985). Agroindustrial wastes such as sugarcane bagasse, wheat bran, rice bran, corncob, and wheat straw were used for xylanase and amylase production from thermophilic actinomycetes Streptomyces sp. The amylase activity (373.89 IU/mL) and xylanase activity (30.15 IU/mL) were maximum in rice bran (Singh et al., 2012).

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4. CONCLUSION Enormous amounts of agricultural wastes are produced worldwide annually. Henceforth, ecofriendly, alternative methods for biowaste handling are needed. Utilization of agricultural wastes as substrates for producing industrially pivotal products such as enzymes, polysaccharides, organic acids, aroma, and flavor compounds by growing microorganisms on these wastes is a valuable technique, with a great economical advantage. Among microbial community, actinobacteria are considered as the potential group for commercial production of various products from wastes since it has unique features and revealed greatest adaptability to extreme environmental conditions. It is anticipated that in the near future industrial bioprocesses will be developed for the production of industrially relevant products.

ACKNOWLEDGMENTS This chapter was supported by research funds of Pukyong National University in 2015. J.S. grateful to University Grants Commission—Dr. D.S. Kothari Postdoctoral Fellowship for financial support.

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CHAPTER SEVEN

Marine Enzymes: Production and Applications for Human Health T. Eswara Rao, M. Imchen, R. Kumavath1 Central University of Kerala, Padannakkad, Kerala, India 1 Corresponding author: e-mail addresses: [email protected]; [email protected]

Contents 1. Introduction 2. Marine Microorganisms 3. Marine Biology and Biotechnology 4. Marine Metagenome as a Resource for Novel Enzymes 5. Applications of Marine Enzymes and Marine Biotechnology for Human Health 6. Conclusion References

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Abstract Marine microbial enzymes have wide applications in bioindustries. Selection of microorganisms for enzyme production at the industrial level requires good yield and high production rate. A number of enzymes such as amylase, caseinase, lipase, gelatinase, and DNases have been discovered from microbes isolated from extreme marine environments. Such enzymes are thermostable, tolerant to a varied range of pH and other harsh conditions required in industrial applications. Novelty in their structure and characteristics has shown promising scope to the researchers in academia and industry. In this chapter, we present a bird’s eye view on recent research works in the field of enzyme production from marine origin as well as their potential biological applications relevant to human health.

1. INTRODUCTION Marine environment is the largest aquatic ecosystem on the planet and most important sources of biodiversity in the world (Zhao, 2011). It houses a wide range of microbes such as archaea, bacteria, fungi, viruses, and protists. The enormous pool of biodiversity in marine ecosystems is a natural reservoir for acquiring an inventory of enzymes with potential novel biocatalysts (Zhang & Kim, 2010) for biotechnological applications. Some microbial Advances in Food and Nutrition Research, Volume 80 ISSN 1043-4526 http://dx.doi.org/10.1016/bs.afnr.2016.11.006

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2017 Elsevier Inc. All rights reserved.

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products based on marine biotechnology are in use in food processing, which are discussed in the later sections. The ever-demanding needs of enzymes in the industrial and pharmaceutical field have tremendously soared up in the past few years, which are expected to increase in the coming future. Metagenomic, a culture-independent study of microbial communities, has the potential to screen novel enzymes in the microbial communities from the environment without having to culture. Hence, metagenomics has provided the scientific community with a variety of novel enzymes (Zhang & Kim, 2010). Microbial enzymes, as opposed to chemical catalysts, enhance the bioprocess reactions economically and eco-friendly through their special characteristics such as tolerance to a wide range of pH, temperatures, and other harsh reaction conditions. Such enzymes are already being utilized in various industries such as food, leather, textiles, and animal feed, and in bioconversions and bioremediations (Nigam, 2013). Marine enzyme displays an effective array of pharmacological applications including pigments, enzyme inhibitors, pesticides, herbicides, toxins, antiparasitics, mycotoxins, antitumor agents, antibiotics, cytotoxicity activities, and growth promoters of animals and plants (Hamdache et al., 2011).

2. MARINE MICROORGANISMS The marine ecosystem covers about 70% of the earth surface. Marine environment provides an extremely complex and diverse natural environment sustaining a wide range of microbial population that exhibits seasonal and spatial variation (Oberbeckmann et al., 2014). The rich microbial diversity in the oceans provides a vast treasure of biological resource, which can be harnessed for wide applications. Microorganisms are the most abundant organisms in the oceans. Marine organisms include 178,000 species falling under 34 phyla (UEPA, 2006). The marine microbial population is largely unexplored such as marine protists being the least explored (Sogin et al., 2006) and rare taxas known as “rare biosphere” being discovered through high-throughput technologies (Caron et al., 2009). Such new technologies have shown the presence of approximately 20,000 species per liter of marine water samples (Sogin et al., 2006). Marine microorganisms are increasingly being studied and becoming a hot spot in the search for industrially important biomolecules. It is estimated that the biological diversity in marine ecosystems is much higher than in the tropical rain forests (Haefner, 2003). The selective force rendered to the microorganisms by various abiotic factors

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such as temperature, altitude, salinity, nutrient availability, and its surrounding environment forces the need to develop multifarious enzymes to adapt to the harsh conditions. Microorganisms found in such environments are potential sources of novel (Hong et al., 2009; Solanki, Khanna, & Lal, 2008) and structurally complex bioactive molecules (Ramesh & Mathivanan, 2009). The potential to discover industrially important biomolecules has attracted researchers from academia and industry (Manivasagan et al., 2013, 2015). Bacteria account for 70% of the overall bioactive compounds discovered from marine sources, while fungi and other domains account for the rest (Prabavathy, Mathivanan, & Murugesan, 2006). Actinobacteria are the most important group of organisms being studied extensively for the discovery of drugs and other bioactive metabolites (Das, Ward, & Burke, 2010). Marine microorganisms are a boundless source for important novel compounds having antagonistic activity against pathogenic organisms.

3. MARINE BIOLOGY AND BIOTECHNOLOGY Marine ecosystem is a vast resource of industrial and pharmaceutical compounds. Besides microorganisms such as bacteria, fungi, and actinobacteria, many other marine organisms such as fishes, prawns, crabs, snakes, plants, and algae have been studied to tap the arsenal of the marine world. Marine microorganisms in extreme environmental conditions such as low temperature, high salinity, and extreme pressures have evolved special metabolites to survive, proliferate, and facilitate storage, transport, and turnover of key biological elements. Marine ecosystems exhibit interesting characteristics that result from the unique combination of several physical factors. These habitats allow the growth of organisms, i.e., bacteria, algae, sponges, fungi, and fishes, which are able to face these harsh conditions. Certain microorganisms are able to live in the cold sea of Arctic and Antarctic regions exerting under high pressures, low temperatures, different pHs, and salinity (Faulkner, 2001). Many of these microorganisms are used in a wide range of biotechnological applications, providing novel bioactive compounds and biocatalysts for modern industries (De Pascale, De Santi, Fu, & Landfald, 2012; Kennedy et al., 2011). Selected microorganisms have been characterized, designed, and optimized to produce high-quality enzyme preparation on large scales for different industrial applications. Recent molecular biology techniques have allowed to tailor specific microorganism to produce not only the high

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yields of desirable enzyme but also the enzyme with desired special characteristics such as thermostability, tolerance at high temperature, and increased stability in acidic or alkaline environment, and to retain the enzyme activity under severe reaction conditions such as the presence of other metals and compounds. The correct in-depth understanding of the properties and functions of genome is a fundamental task in modern bioscience. Molecular biology has a major role in many aspects of marine biotechnology. Genome study of different marine microorganisms can facilitate the use of genes in cell factories and bioindicator strains as well as the identification of new drug targets. A flowchart on the use of omics methodologies for the identification of novel biocatalysts from marine ecosystems is shown in Fig. 1. Marine sponge is a primitive organism in animal kingdom and is described as a living fossil. Genome analysis of primitive organism is a special interest in molecular evolution. Applications of molecular biological techniques have proved to be beneficial to Marine genome projects, which were performed until January 2009 (Fig. 2). The values include complete and incomplete genome sequencing projects based on the information by Genomes Online.

Microbial biomes from marine ecosystem

Extraction of protein

Genomic DNA from marine microbes

Metaproteomic analysis by 2D-PAGE or tandem MS

Construction of metagenomic library

Sequence-derived analysis

Function-derived analysis

Cloning and heterologous expression

Gene identification by probes

Gene identification by bioinformatics analysis Gene analysis/cloning and expression

Screening regimes Identification of products

Fig. 1 Omics-based approaches to identify novel biocatalysts from marine ecosystem.

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Marine genome project

10 (2.7%)

38 (10.0%) Bacteria Archaea Eukarya

331 (87.3%) Current Opinion in Biotechnology

Fig. 2 Scheme of the Marine genome project. Adapted from Lee, H. S., et al. (2010). Approaches for novel enzyme discovery from marine environments. Current Opinion in Biotechnology, 21(3), 353–357.

4. MARINE METAGENOME AS A RESOURCE FOR NOVEL ENZYMES More than 98% of the microorganisms cannot be cultivated yet in the laboratory. It can be understood that a huge portion of the microbial based bioactive compounds and enzymes are being missed out by the traditional culture-based technique. Metagenomics is the study and analysis of microbial communities from the environment without culturing. It also provides the potential to discover novel enzymes through functional-based screening. Thus, metagenomic is a powerful tool for the exploitation of bioactive compounds from marine microbial communities such as sponges, which has been indicated as a promising source of novel compounds by an array of literature (Kennedy, Marchesi, & Dobson, 2007; Schirmer et al., 2005). Metagenomic libraries constructed in different hosts and vectors have the ability to enrich the current toolbox of enzymes used in industry, therapeutics, and environmental sustainability ( Kennedy, Marchesi, & Dobson, 2008). The advances in metagenomics have revolutionized the research in fields of microbial ecology and biotechnology, enabling not only a glimpse into the uncultured microbial population and mechanistic

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understanding of possible biogeochemical cycles and lifestyles (Zhao, Chen, Chen, Wang, & Geng, 2015) of extreme organisms but also the highthroughput discovery of new enzymes for industrial bioconversions.

5. APPLICATIONS OF MARINE ENZYMES AND MARINE BIOTECHNOLOGY FOR HUMAN HEALTH Microbial enzymes are in higher demand compared to the animal and plant sources largely because of the high growth rate, minimal space requirements, inexpensive media, ethics, and the availability of enzymes with catalytic activity in a wide array of environments. The large amount of such stable enzymes shows a great deal of biological activities such as antibacterial, antifungal, anticancer, antitumor, antiinflammatory, antimalarial, antiviral, cytotoxic, and antiangiogenesis drugs (Table 1). The marine microbial world has evolved through a series of various environmental conditions. Thus, the marine ecosystem may harbor unique species with potential novel antimicrobial agents (Kusaykin et al., 2003). Hence, efforts should be made to make the best use of the rich biological diversity. Among the enzymes listed in Table 1, proteases and lipases have several applications for human health. So, proteases have a wide array of applications in pharmaceuticals such as digestive drugs and antiinflammatory drugs (Kumar et al., 2004). In the last decade, a number of proteases have been isolated from various sources with different properties (Haddar et al., 2009). Microbial lipases were isolated from Penicillium oxalicum, Aspergillus flavus, and Streptomyces (Mo, Kim, & Cho, 2009). These enzymes can actively hydrolyze different oils and organic solvents and show a broad substrate specificity (Wang et al., 2007). Many of the lipases currently in use are spread in the production of pharmaceuticals, food flavorings, and other industrial applications (Basheer et al., 2011). Marine natural products play an important role in biomedical research and drug development (Debbab, Aly, Lin, & Proksch, 2010). The need to explore the untapped resource in the various marine environments has been realized through the novel discoveries. Studying the microbial abundance and diversity would lead to a better understanding in the various biogeochemical cycles and realization of the rich source of microbial bioactive molecules. Omics-based studies involving high-throughput sequencing and screening have the potential to solve such issue.

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Table 1 Stable Enzymes Show a Great Deal of Biological Activities Enzyme Name Source of Bacteria Function

References

Hwang et al. (1997)

Alpha amylase (3.2.1.1)

Bacillus licheniformis

Hydrolysis of alpha 1,4 bonds passing alpha 1,6 bonds (endoaction)

Protease (EC 3.4.21.88)

Bacillus, Pseudomonas, Clostridium, Rhizopus, Penicillium, Aspergillus

Nigam and Acidic neutral alkaline thermophilic Pandey (2009) active in the presence of inhibitory compounds

Xylanase (EC 3.2.1.8)

Thermoactinomyces thalophilus: Humicola insolens, Bispora (acidophilic fungus)

Extremophilic characteristic— alkalophilic thermophilic and thermostable

Ligninase (EC 3.2.1.8)

Basidiomycetes

Lignin hydrolysis

Cellulose (232-674-9)

Cellulose hydrolysis Basidiomycetes strains, Polyporus sp., Pleurotus sp., Trichoderma sp., Aspergillus sp.

Nigam and Prabhu (1988)

Lipase (EC 3.1.1.4)

Yeast and fungi

Lipid hydrolysis

Muralidhar et al. (2002)

Keratinase (EC 3.4.99)

Actinomycetes, fungi

Keratin hydrolysis

Gupta, Sharma, and Beg (2013)

Amylase (EC 3.2.1.1)

Bacillus sp., Geobacillus

Starch hydrolysis

Singh, Dahiya, and Nigam (1995)

Beta-galactosidase Marine mollusc (EC 3.2.1.23)

Alphagalactosidase (EC 3.2.1.33)

Arthrobacter sp., SB

Hydrolysis of O-β-D- Giordano, galactopyranosides Andreotti, Mollo, and Trincone (2004) Joint, M€ uhling, Hydrolyzes the and Querellou terminal alphagalactosyl moieties (2010) from glycolipids and glycoproteins Continued

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Table 1 Stable Enzymes Show a Great Deal of Biological Activities—cont’d Enzyme Name Source of Bacteria Function References

Epoxide hydrolase Rhodobacterales (EC 3.3.2.10) bacterium

Woo et al. Highly (2013) enantioselective toward racemic glycidyl phenyl ether

β-amylase (EC 3.2.1.2)

Hydrolysis of alpha 1,4 bonds passing alpha 1,6 bonds (exoaction)

Balls, Walden, and Thompson (1948)

β-D-mannosidases Marine Anaspidea, (EC 3.2.1.25) Aplysia fasciata

The betamannopyranoside linkage

Andreotti, Giordano, Tramice, Mollo, and Trincone (2005)

Superoxide dismutase (EC 1.15.1.1)

Antioxidant genes

McCord and Fridovich (1969)

Alpha-L-fucosidase Marine mollusk (EC 3.2.1.51)

Hydrolysis of L-fucoidan

Berteau et al. (2002)

Esterase (EC 3.1.1.1)

Seashore sediments

Antibiotic resistance

Jeon et al. (2009a)

Lipase (EC 3.1.1.4)

Deep sea

Lipids hydrolysis

Jeon et al. (2009b)

Esterase (EC 3.1.1.85)

Interdial zone

Antibiotic resistance

Fu et al. (2013)

Alkaline phospholipase (EC 3.4.21.67)

Tidal flat sediments

Hydrolysis

Lee et al. (2010)

Glycoside (EC 3.2.1)

Baltic sea

Hydrolysis

WierzbickaWos´ et al. (2013)

Chloroperoxidase (EC 1.11.1.18)

Caldariomyces fumago Halogenation reactions

Morris and Hager (1966)

Phospholipase (EC 3.1.4.11)

Hot spring

Hydrolysis

Tirawongsaroj et al. (2008)

Esterase (EC 3.1.1.1)

Deep-sea hydrothermal field

Antibiotic resistance

Zhu et al. (2013)

Bacteria, fungi

Cyanobacterium, Synechococcus sp.

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Table 1 Stable Enzymes Show a Great Deal of Biological Activities—cont’d Enzyme Name Source of Bacteria Function References

Glycoside hydrolases (EC 3.2.1)

Hydrothermal vent

Degradation

Wang et al. (2011)

Fumarase (EC 4.2.1.2)

Marine water

Hydrolysis

Jiang et al. (2010)

Beta-glucosidase (EC 3.2.1.21)

Hydrothermal spring Hydrolysis

Rath and Herndl (1994)

Laccase (EC 1.10.3.2)

Marine water

Oxidation

Fang et al. (2012)

Esterase (salt tolerant) (EC 3.1.1.1)

Tidal flat sediment

Novel phenotypic on Jeon et al. host organism (2012)

Esterase (EC 3.1.1.1)

Red sea brine pool

Hydrolysis

Mohamed et al. (2013)

Mercuric reductase (EC 1.16.1.1)

Red sea brine pool

Reduction process

Sayed et al. (2014)

Glucoamylase (EC 3.2.1.3)

Aspergillus oryzae

Hydrolysis of 1,4 and Hata et al. 1,6 bonds (1997)

Pullulanase (EC 3.2.1.41)

Bacillus deramificans

Hydrolysis of alpha 1,6 bonds

Marine water Cyclodextrin glycosyltransferase (EC 2.4.1.19)

Hii, Tan, Ling, and Ariff (2012)

Hydrolysis of starch Ritter (2012) to nonreducing cyclic D-glucose polymers Degradation of agar- Parro et al. degrading bacteria an (1999) abundant food source ocean

Agarases (EC 3.2.1.81)

Cytophaga, Bacillus, Vibrio, Alteromonas, Pseudoalteromonas, Streptomyces

Cellulase (EC 3.2.1.4)

Cytophaga, Cellulose hydrolysis Cellulomonas, Vibrio, Clostridium, Nocardia, Streptomyces

Maki, Leung, and Qin (2009)

Chitinase and chitosanase (EC 3.2.1.14)

Degradation Aspergillus, Penicillium, Rhizopus, Myxobacter, Sporocytophaga, Bacillus

Jolle`s and Muzzarelli (1999)

Continued

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Table 1 Stable Enzymes Show a Great Deal of Biological Activities—cont’d Enzyme Name Source of Bacteria Function References

Gelatinase (EC 3.4.24.24)

Marine water

Hydrolysis

Hibbs, Hoidal, and Kang (1987)

Lipase (EC 3.1.1.4)

Moraxella

Lipid hydrolysis

Svendsen (2000)

Laccase/ligninase (EC 1.10.3.2)

Plants, fungi

Oxidation

Selinheimo et al. (2007)

Luciferase (EC 1.13.12.7)

Photinus pyralis

Oxidation

Gould and Subramani (1988)

Xylanase (EC 3.2.1.8)

Marine algae

Xylan hydrolysis

Subramaniyan and Prema (2002)

Protease (EC 3.4.21.40)

Bacillus licheniformis

Protein hydrolysis

Oda (2012)

Marino pyrroles A (EC 1.2.2.2)

Marine bacteria

Antibacterial

Trisuwan et al. (2009)

Marino pyrroles B (EC 1.2.2.3)

Marine bacteria

Antimicrobial

Butler and Buss (2006)

Nigrospoxydons Marine fungi and A–C (EC 1.11.1.5) bacteria

Antimicrobial

Fremlin, Piggott, Lacey, and Capon (2009)

Haloperoxidase (EC 1.11.1.10)

Marine sponge— Druinella purpurea

Oxidation of halides

Butler and Walker (1993)

Collagenase (EC 3.4.24.3)

Clostridium histolyticum

Cleaving native collagen types

Mandl (1981)

Superoxide dismutase (EC 1.15.1.1)

LE392 E. coli

Superoxide (O2 ) radical into either ordinary molecular oxygen (O2)

Mark D. Scot

DNA ligase (EC 6.5.1.1)

Thermococcales

Ligation

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6. CONCLUSION The need for specialized and active enzymes in the market is in demand. Oceans represent 70% of the earth’s surface, which harbors more species than a rainforest. Marine ecosystem harbors an enormous array of enzymes with potential for industrial and pharmaceutical applications. Various proteases, lipases, antioxidants, and a wide array of enzymes have been discovered in the recent past from marine sources. However, conventional culture-based methods are unable to harness their potential. A combination of marine biotechnology and metagenomics with high-throughput screening techniques is the need of the hour to harness the unseen part of the marine ecosystem.

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CHAPTER EIGHT

Bioremediation of Industrial Waste Through Enzyme Producing Marine Microorganisms P. Sivaperumal*,1, K. Kamala*, R. Rajaram† *Center for Environmental Nuclear Research, Directorate of Research, SRM University, Kattankulathur, India † Bharathidasan University, Tiruchirappalli, India 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Role of Microorganisms and Their Enzymes in Marine Environment 3. Industrial Waste Around Marine Environment 4. Bioremediation of Industrial Waste 5. Marine Enzymes: Decontaminating Agents 6. Conclusion Acknowledgments References

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Abstract Bioremediation process using microorganisms is a kind of nature-friendly and costeffective clean green technology. Recently, biodegradation of industrial wastes using enzymes from marine microorganisms has been reported worldwide. The prospectus research activity in remediation area would contribute toward the development of advanced bioprocess technology. To minimize industrial wastes, marine enzymes could constitute a novel alternative in terms of waste treatment. Nowadays, the evidence on the mechanisms of bioremediation-related enzymes from marine microorganisms has been extensively studied. This review also will provide information about enzymes from various marine microorganisms and their complexity in the biodegradation of comprehensive range of industrial wastes.

1. INTRODUCTION Enzymes derived from marine microorganisms have more advantages when compared with other derived enzymes from plants or animals. It has various catalyzing activity, solvent and temperature stability, high Advances in Food and Nutrition Research, Volume 80 ISSN 1043-4526 http://dx.doi.org/10.1016/bs.afnr.2016.10.006

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production in less time, and is easy to harvest. In older days, enzymes from terrestrial sources have been used for many industrial applications with fewer outcomes. However, enormous sources in marine-derived enzymes and their exploration are still limited (Chandrasekaran, 1996). Marine microbial enzymes are explored from various marine sources including mangrove, seagrass, coral reef, open sea, deep sea, coastal region, estuarine, brackish water, lagoon, and hydrothermal vent samples. In addition, marine microorganisms including archaea, bacteria (actinobacteria), fungi, viruses, and many other marine organisms were also studied to tap the resource of the marine world (Sarkar, Roy, & Mukherjee, 2010). Such enzymes may have various industrial applications in the form of protease, amylase, and lipase enzymes (Kumar & Takagi, 1999). Out of three prime industrial enzymes, proteases take maximum 60% of contribution in universal market (Rao, Tanksale, Ghatge, & Deshpande, 1998). In addition, enzyme-facilitated practices are promptly gaining attention due to the less process time; low energy input; and less toxic, cost-effective, and ecofriendly features (Choi, Han, & Kim, 2015; Li, Yang, Yang, et al., 2012). Moreover, with the advanced technology of recombinant DNA techniques and microbial protein engineering could be influenced and microbial cultured in huge quantities to encounter the increased demand (Liu, Yang, & Shin, 2013). Marine enzymes are exactly converting the complex material of polysaccharide (exo and endo) secreted by the microbial communities in the marine environment. Complex materials like polysaccharides are well known for bioremediation of various pollutants in water and sediment. During the bioremediation, microorganisms secrete the polysaccharides which bind with pollutants and settle them down as deposition; simultaneously deposition was used as the source of nutrients. So, the process will reduce the rate and risk of contaminants or pollutants, yet the process was relaxable (Karigar & Rao, 2011). However, microbial growth greatly depends on physical, chemical, biological, and nutrient availability in different environmental factors such as temperature, salinity, pH, BOD, COD, oxygen, moisture, organic nutrients, and occurrence of supplementary toxic mixtures and other extreme environment conditions (Bernhard-Reversat & Schwartz, 1997; Dana & Bauder, 2011; Vidali, 2001). Some rare marine species of archaea, fungi, and bacteria have capacity to swift the complete bioremediation process. But in laboratory conditions, only some microbial strains are working effectively as bioremediation mediators (Kumar, Bisht, Joshi, & Dhewa, 2011). Therefore, this chapter is focused on marine

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microbial enzyme and their potential application in industrial waste water treatment.

2. ROLE OF MICROORGANISMS AND THEIR ENZYMES IN MARINE ENVIRONMENT Over and above 3.2 billion years, microbial life on earth and their biogeochemical cycle were alive. Biological system of the environment was improved by the oxygen uptake from well-developed microorganisms in the formation of primary production, which created macroscopic inhabitation in the earth. According to environmental conditions on earth, the microorganisms have been also involved in continuous changes and get adapted to global environmental changes. Generally, the distinction between terrestrial and aquatic microorganisms is relatively clear, yet scientific approaches between aquatic and soil microorganisms differ. This hypothetical difference derives from salinity which is an important factor for microbial growth. Similarly, fresh water microorganisms could not survive within the range of salinity from 30 to 35 ppt, because osmotic regulations of the microbial cells in the marine and fresh water environments are entirely contradictory to each other. Very few microorganisms can survive with broad series of environmental parameters such as temperature, salinity, and pH (Glockner et al., 2012). In addition, microorganisms also grow in marine environs including low-salinity area such as brackish water or estuaries, moderate saline environments, deep ocean brines, and solar salt environment. Marine microorganisms have immense genetic and biochemical diversity; therefore, researchers are more interested in promising new enzymes and other secondary metabolites exploration and their hyperthermostability, salt tolerance, and cold adaptation properties. Marine microorganisms are essential to major biogeochemical cycles, changes, and processes occurring in marine systems and are useful in the movement between reduced and oxidized forms. Therefore, marine microorganisms are critically important to environmental as well as human health. They are tremendously abundant and diverse, playing a vital key role in the regulation of Earth’s climate as it can release carbon products, particularly CO2 and CH4. They also provided essential goods and services to humans in terms of oxygen production, supplying sustainable food, regulating marine environment for unexplored genetic information and significant biomolecules. These marine microorganisms are capable to transfer

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C-, N-, P-, and S-containing compounds for biological production, and finally it will influence their availability for natural sources. These metabolisms in marine microorganisms maintain significant oxygen production required for aerobic life and carbon for biological removal and biogeochemical cycles on Earth. These biogeocycles are stabilizing the entire ocean biosphere; therefore, special distribution of marine microbial studies gets focused in all ecosystems. The waste management system enzymes are using widely different kinds of enzyme complexes in remediation of toxic pollutants. The domestic waste and industrial sewages are having many chemical types of merchandise, which are toxic to the ecosystem as well as human being. To minimize these hazardous materials, microbial enzymes, in combinations (two or more enzymes together) or alone, were used for the management of industrial waste containing nitriles, phenols, and aromatic amines by the degradation of noxious chemical compounds to harmless products (Pandey, Singh, & Chand, 2011; Rubilar, Diez, & Gianfreda, 2008). A number of enzymes are functioning for waste treatment such as amylases, amyloglucosidases, amidases, glucoamylases, cellulases, lipases, proteases, and pectinases (Karigar & Rao, 2011; Riffaldi, LeviMinzi, Cardelli, et al., 2006). One of the major enzymes, protease is frequently isolated from marine microorganisms and shows essential function in the detergent preparing industry, tanneries, and pharmaceutical industries. Likewise, agarase, amylase, cellulase, carragenases, chitinase, lipase, and lignocellulose are also isolated from marine microorganisms and are used in the production of bioethanol and other purposes. Moreover, cellulose chains are typically more complicated and difficult to hydrolyze so that cellulose molecules from marine derivatives might give lower hydrolysis yields. Therefore, the degree of polymerization of cellulose is recognized to be a serious issue when being hydrolyzed (Karimi, Shafiei, & Kumar, 2013). Earlier, Gao et al. (2014) also reported the strong track relation between maximum adsorption capacity and the enzymatic hydrolysis. Oxidoreductases enzymes, like manganese peroxidase, laccase, lignin peroxidase, and tyrosinase, catalyze or eliminate the industrial effluents of chlorinated phenolic compounds (Le Roes-Hill & Prins, 2016). Recently, Karimi and Taherzadeh (2016) critically reviewed the pretreatment of lignocelluloses and changes of the different properties during the adsorption/desorption and accessibility of the enzymes, and also reported enzyme measurement techniques and modification techniques for bioconversions.

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3. INDUSTRIAL WASTE AROUND MARINE ENVIRONMENT The quantity of industrial waste is closely linked to the level of economic activity in a country. A huge variety of hazardous sediment contaminants, with different effects on the marine environment and human health, are also available depending on their possible scattering, solubility in seawater, leakage of oil from container, and carcinogenicity among other characteristics. Mining activities for oils constitute the key class of organic sediment contaminants in marine environment. In addition, the harmful effects of accidental spills on the marine environment depend on the category of oil as well as on the additives, which are significantly involved in toxicity of the spill (Aluyor & Ori-Jesu, 2009). One of the major hazardous materials of chlorinated hydrocarbons used as solvents, preservative agents, pesticides, heavy metals, dyes, and pharmaceutical pollutants is distributed in the marine environment and can cause extreme range of cancers to humans (Badawi, Cavalieri, & Rogan, 2000). A further group of pollutant polycyclic aromatic hydrocarbons are increased by accidental spills, coal, burning, and waste disposal to marine environ and wood preservation. It is recognized to create also lethal effect and carcinogenic activity (Cerniglia, 1993). In addition, ethyl benzene; benzene; toluene; and o-, m-, and p-xylenes are widely used in industrial synthesis and are also dumped in marine environment, leading to carcinogenic and neurotoxic diseases (Levin, Viale, & Forchiassin, 2003). Organophosphorus compounds used as petroleum additives, pesticides, and band plasticizers are involved in many degenerative syndromes of the nervous system and are strong mutagens, causing carcinogenesis and chromosomal aberrations (Singh, 2009; Sirotkina, Lyagin, & Efremenko, 2012).

4. BIOREMEDIATION OF INDUSTRIAL WASTE Earlier, wastes were conventionally disposed by burrowing the waste materials. This traditional method was complicated to bear, because every time there is short for digging ground. New tools of high-temperature incineration and chemical decomposition practices are evolved for waste disposal. These wastes could be very active in decreasing an extensive range of contaminants; nevertheless at the same time they have a number of disadvantages. These traditional methods are difficult, profligate, and lack

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public recognition. The associated scarcities in these methods have motivated to near harnessing modern-day bioremediation technology as an applicable substitute. However, bioremediation using microorganisms is an economical preference, and it has the capacity to degrade the higher-level contaminants into nonhazardous or less hazardous substances. Previous studies suggested that microbial bioremediation was very effective (BernhardReversat & Schwartz, 1997; Leung, 2004). Most bioremediation functions are under aerobic conditions, although degradation of recalcitrant molecules by microorganisms can occur under anaerobic condition also. Altered intracellular and extracellular enzymes from marine bacteria and fungi have participated in the remediation of lignin and organic pollutants (BernhardReversat & Schwartz, 1997; Hammel, 1997). Marine environment receives higher-level pollutants from metal smelting industrial waste, petrochemical industrial waste, mining activities, paper and pulp industry, chemical weapon producing industry, dye industry, anthropogenic activity, and agriculture wastes. Particularly, dye remains in industrial wastewaters constitute an important source of water pollution. Unused dyes, approximately 10–15%, are passed into the wastewater after dyeing and after the subsequent washing processes (Rajamohan & Karthikeyan, 2006). Subsequently water is the greatest vital natural resource; thus treatment of these wastewaters is an unavoidable responsibility. The colored wastewater was treated using many physical and chemical processes, but these skills are unsuccessful in removing dyes in addition to the high cost incurred in the treatment. The biosorption was later recognized as the preferred skill for bleaching of colored wastewater (Rana & Samir, 2014). Ultimately the efficient treatments for color removal from colored effluents comprised of combined approaches of three combined methods such as physical, chemical, and biological for dye removal (Azbar et al., 2004; Galindo & Kalt, 1999; Robinson, McMullan, Marchant, & Nigam, 2001). Generally, physical and chemical treatments are not used in the remediation of textile waste water due to the high cost and nature of the waste. However, the dye effluents are easily remediated by biological organisms of bacterial and fungal isolates (Fu & Viraraghavan, 2002; Yang, Zhao, Liu, Zheng, & Qian, 2009), and they were low-cost nonconventional adsorbents (Crini, 2006; Ferrero, 2007). Tony et al. (2013) studied that white-rot fungus Pleurotus eryngii could decolorize the reactive black 5 azo dye from dye effluent. Similarly, Abedin (2008) also reported crystal violet and malachite green dye removal by Fusarium solani fungus. In addition, Tak et al. (2004) found that high negative charged reactive dye solutions were

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decreased due to white-rot fungi at high pH level. This might be due to the adverse effect of alkaline range of pH in dye biodegradation by fungal enzymes. The decontamination of toxic organic compounds through oxidation coupled with oxidoreductases using different bacteria and fungi (Gianfreda, Xu, & Bollag, 1999) and higher plants was also examined (Bollag & Dec, 1998). These microbial enzymes are intermediate in the energy-yielding biochemical reaction by electron exchange from substrate which leads to cleavage in chemical bonds. During the oxidation and reduction process, the organic pollutants were converted into nontoxic or simple compounds (Interstate Technology and Regulatory Council [ITRC], 2002). Moreover, the enzyme oxidoreductase is able to decompose the lignin by humified phenolic substances and detoxify the aniline-based xenobiotics during the course of polymerization and copolymerization binding with humic substances (Park, Park, & Kim, 2006). In addition to that, microbial enzymes are significantly able to degrade and decolorize the azo dyes obtained from industrial wastes (Bernhard-Reversat & Schwartz, 1997; Husain, 2006; Williams, 1977). Similarly, radioactive substances are also remediated using diverse microorganisms by the oxidation process, and these microorganisms can convert radioactive substances from soluble to insoluble stage. During this oxidation process microorganisms utilize the organic substances to provide the electrons for redox reaction which leads to precipitation of insoluble radioactive metal (Leung, 2004). Partially degraded lignin and chlorinated phenolic compounds are sources of pollutants released into the environment from paper and pulp industries. These wastes are bioremediated with fungal species that produce extracellular enzymes (oxidoreductase), viz., manganese peroxidases, lignin peroxidases, and laccase from fungal mycelium, being a quicker bioremediation process than bacteria (Rubilar et al., 2008). Usually, oxygenase enzymes are grouped into two types such as mono- and dioxygenases, on the basis of oxygen atoms utilization in oxygenation process. Oxygenases have a key position to develop water solubility and lead to cleavage of aromatic ring in a variety of chlorinated aliphatics and organic pollutants. Above all, the oxygenase from marine bacteria could be a mono- or dioxygenase, and some studies have reported the role of oxygenases enzymes in the process of biodegradation of organic pollutants (Arora, Kumar, Chauhan, Raghava, & Jain, 2009; Fetzner, 2003; Fetzner & Lingens, 1994). Marine-derived fungal strains of Aspergillus niger and Cerrena unicolor also produced alkaline xylanases and thermostable metal-tolerant

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laccases (D’Souza-Ticlo, Sharma, & Raghukumar, 2009; Raghukumar, Muraleedharan, Gaud, & Mishra, 2004) while degrading the pollutant. Many fungal strains are known to be capable of degrading persistent pollutants including textiles dyes (Haritash & Kaushik, 2009). Passarini, Rodrigues, DaSilva, and Sette (2011) isolated salt-tolerant fungi and their lignin-degrading enzymes for environmental pollutants bioremediation. The dye decolorization process by fungal cells includes oxidative reactions, which could produce nontoxic derivatives (Ciullini, Tilli, Scozzafava, & Briganti, 2008). Marine filamentous fungal extracellular enzymes of lignin lytic will be given excessive importance in environmental remediation (Arun et al., 2008). In addition to that, other research groups also have been concentrated on the use of marine-derived filamentous fungi for man-made dye decolorization (Bonugli-Santos, Durrant, & Sette, 2012; Chen, Wang, Shen, & Yao, 2014; Junghanns, Krauss, & Schlosser, 2008), and earlier Chen et al. (2014) reported that marine-derived fungal Pestalotiopsis sp. J63 and Penicillium janthinellum P1 cell immobilization system could remove the Azure B.

5. MARINE ENZYMES: DECONTAMINATING AGENTS Enzymes have a number of applications in different fields, and so they can virtually catalyze a mixture of diverse pollutants from level of soluble toxic to insoluble nontoxic materials. This microbial bioprocess is much effective and there are no harmful chemical alterations. In addition, enzymes have more advantages and traditional technologies as well as cover microbial remediation process. Certainly, enzymes are not inhibited by microbial metabolic inhibitors. They are effective at low-level contaminants that are treated with microbial antagonistic properties. Generally, enzymes are more active than microorganisms to utilize the substrate and its transfer from complex to simple form or smaller substances (Gianfreda & Bollag, 2002). Most of the enzymes are ecologically beneficial and risk-free for the environment, being able to play a vital role in the bioremediation of adverse compounds in the environment. The enzymatic catalysis can be considered as green chemistry (Alcalde, Ferrer, Plou, & Ballesteros, 2006; Sheldon & Van Rantwijk, 2004). In addition, immobilization of biomolecules and the role of enzymes in bioremediation of pollutants are illustrated (Rodrı´guez Couto & Toca Herrera, 2006) by laccase, an enzyme frequently used in decontamination of pollutants (Gianfreda et al., 1999).

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Hydrolases are amidases, proteases, carbohydrases, phosphotriesterases, and depolymerases. The main classes of oxidoreductases are mono- or dioxygenases, dehalogenases, reductases, cytochrome P-450 monooxygenases, peroxidases, and phenoloxidases. In addition to that, bacterial hydrolases of parathion or carbamate from Pseudomonas, Flavobacterium, Achromobacter, Nocardia, and Bacillus cereus have effectively degraded the diazinon, coumaphos, parathion, and carbofuran. But sometimes protases, amidases, and esterases cause the disruption of amidic, peptidic, and esteric bond formation, leading to the production of toxic or nontoxic products (Coppella et al., 1990; Mulbry & Eaton, 1991; Sutherland, Russel, & Selleck, 2002). Likewise, depolymerases, carbohydrases, phosphatases, and proteases from marine microorganisms convert the insoluble material like proteins, carbohydrates, and plastics into soluble harmless form (Nakamura, Tomita, Abe, & Kamio, 2001; Singh, 2002; Van Wyk, 1999). Recently, Vasconcelos et al. (2015) reported that marine-derived fungi produced hydrolytic enzyme and oxidative enzymes such as amylase, cellulase, chitinase, lipase, glucosidase, keratinase, ligninase, protease, and xylanase. All these enzymes are highly active at temperatures ranging from 35°C to 70°C and pH ranging from 3.0 to 11.0. The creation of industrially related enzymes from marine environment genomes has novelty and potential applications, supported by advanced technologies such as metagenomics and culturing (Drepper et al., 2014). In addition, it is significant to increase the knowledge on effective enzyme selection protocol and sequence-based description for worldwide marine environments through metagenomic tools (Feller, 2013), which may have promising novel biotechnological applications (Niehaus et al., 2011), in extreme conditions (Alcaide et al., 2015; Ferrer et al., 2016; Vester, Glaring, & Stougaard, 2014). In addition, biocatalysts both in the form of immobilized or free enzyme, cell-free system and complete cell catalysts was highly applicable for the remediation process (Jeon, Baek, Bornscheuer, & Park, 2015; Schmidt et al., 2015; Schrewe, Julsing, B€ uhler, & Schmid, 2013; You & Zhang, 2013). The most relevant enzymes for the detoxification of organic pollutants are oxidative enzymes (Duran & Esposito, 2000; Gianfreda, Iamarino, Scelza, & Rao, 2006; Gianfreda & Rao, 2004; Gianfreda et al., 1999; Rodrı´guez Couto & Toca Herrera, 2006; Torres, Bustos-Jaimes, & Le Borgne, 2003). For instance, nonspecific oxidative lignolytic enzyme is involved in dye degradation and minimizes the lignolytic complex and many other environmental pollutants which could not be degraded with any other microorganisms (Pointing, 2001; Reddy, 1995). These lignolytic enzymes have primary relevance in various

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environmental bioremediation processes. A large amount of this enzyme is produced by white-rot fungi and is utilized to remediate the pollutants present in the environment at high concentration. It is more effective than bacterial oxidative enzymes (Asgher, Nawaz Bhatti, Ashraf, & Legge, 2008; Bumpus, 1993; Rubilar et al., 2008).

6. CONCLUSION In conclusion, there are many fields in which marine microbial enzymes can be applied for industrial waste treatment. The aim of bioremediation using marine microbial enzymes is to minimize the amount of pollutants present in the environment, reducing them to negligible level with low cost and in a safe mode. In addition, the marine microbial remediation is evaluated as a green technology and most effective applicable process for industries to minimize risks related with hazardous wastes. This review provides information on the marine microbial-derived enzymes and their biodegradation of a range of industrial pollutants and their applications. Finally, the uses of enzymes from marine microorganisms are perhaps a more appropriate tool for use against industrial contamination in and around the marine environment.

ACKNOWLEDGMENTS The authors are grateful to Director CIFE, ICAR, Deemed University, Mumbai, and Department of Marine Science, Bharathidasan University, Tamil Nadu, and thankful to the Center for Environmental Nuclear Research, Directorate of Research, SRM University, Tamil Nadu, for providing facilities. The second author would like to thank Science and Engineering Research BoardDepartment of Science and Technology, Govt. of India for providing fund through National Post-Doctoral Fellowship program (File No: PDF/2015/000680).

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CHAPTER NINE

Marine Enzymes and Microorganisms for Bioethanol Production M.R. Swain, V. Natarajan, C. Krishnan1 Indian Institute of Technology Madras, Chennai, India 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Bioethanol Production Technology 3. Marine Enzymes Used for Bioethanol Production 3.1 Production and Biochemical Properties 3.2 Cloning and Characterization 4. Marine Microoorganisms Producing Ethanol 5. Bioethanol Production From the Marine Algae 6. Conclusion Acknowledgments References

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Abstract Bioethanol is a potential alternative fuel to fossil fuels. Bioethanol as a fuel has several economic and environmental benefits. Though bioethanol is produced using starch and sugarcane juice, these materials are in conflict with food availability. To avoid food–fuel conflict, the second-generation bioethanol production by utilizing nonfood lignocellulosic materials has been extensively investigated. However, due to the complexity of lignocellulose architecture, the process is complicated and not economically competitive. The cultivation of lignocellulosic energy crops indirectly affects the food supplies by extensive land use. Marine algae have attracted attention to replace the lignocellulosic feedstock for bioethanol production, since the algae grow fast, do not use land, avoid food–fuel conflict and have several varieties to suit the cultivation environment. The composition of algae is not as complex as lignocellulose due to the absence of lignin, which renders easy hydrolysis of polysaccharides to fermentable sugars. Marine organisms also produce cold-active enzymes for hydrolysis of starch, cellulose, and algal polysaccharides, which can be employed in bioethanol process. Marine microoorganisms are also capable of fermenting sugars under high salt environment. Therefore, marine biocatalysts are promising for development of efficient processes for bioethanol production.

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1. INTRODUCTION Bioethanol has been recommended as an alternate fuel to gasoline due to its economic and environmental benefits. Since the feedstocks for bioethanol are renewable plant materials, there is no net emission of carbon to atmosphere due to the use of bioethanol as a fuel (Arora, Behera, & Kumar, 2015). The plant feedstocks can be categorized into (i) sugar syrups like sugarcane juice, (ii) starchy grains and tubers, and (iii) cellulosic materials such as stems and leaves. At present, starchy grains like corn and sugarcane juice are widely used as raw materials for the production of bioethanol (Dias, Junqueira, Rossell, Filho, & Bonomi, 2013). Since these two sources belong to food category, use of these materials competes with food supplies. The food–fuel conflict can be avoided by using cellulosic feedstocks such as rice straw, wheat straw, sugarcane bagasse, and wood. Though the cellulosic feedstocks are relatively low cost, their conversion into bioethanol is a complex and expensive process (Menon & Rao, 2012). Recently, algal biomass as feedstock has attracted attention for the production of bioethanol. The use of algal biomass has many benefits such as no land use and short growth cycle (Daroch, Geng, & Wang, 2013). Based on the nature of feedstock, bioethanol production can be classified into three generations: (i) first-generation bioethanol from edible materials like corn and sugar cane juice, (ii) second-generation bioethanol from lignocellulosic residues of food crops and energy crops, and (iii) thirdgeneration bioethanol from algal biomass. While first-generation feedstocks have the food vs fuel conflict, the second-generation feedstocks have other issues such as land use and conflict with animal feed (Menon & Rao, 2012). The third-generation feedstocks are marine based such as microand macroalgae, which have an edge over the first- and second-generation feedstocks with respect to conflict with food, feed, and land use. In addition, several marine organisms produce enzyme biocatalysts for saccharification of diverse polysaccharides that are present in the feedstocks used for bioethanol production and possess capability to ferment multiple sugars to ethanol (Ge, Wang, & Mou, 2011). Here, we describe the potential of marine organisms as sources of biocatalysts and feedstock for the production of bioethanol.

2. BIOETHANOL PRODUCTION TECHNOLOGY The overall processes for three generations of bioethanol production are shown in Fig. 1. First-generation bioethanol production uses food-based

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Sugars/ carbohydrates

Biomass

Conversion step

1st generation Biological Sugar crops

Sugars

Grains

Starches

Conventional alcohol fermentation

Enzymatic hydrolysis and fermentation

2nd generation

Ethanol

Agricultural residues Cellulose hemicellulose

Trees and grasses

Algae

Chemical hydrolysis and fermentation

Pretreatment

3rd generation

Chemical

Fig. 1 Outline of ethanol production processes from three different types of feedstocks.

feedstock such as starch and sugar juice. While sugar juice is fermented directly to ethanol, starchy materials like grains require hydrolysis of starch polysaccharide to monomeric glucose. Starch is hydrolyzed to glucose using enzymes α-amylases and glucoamylases (Naik, Goud, Rout, & Dalai, 2010). The starch hydrolyzate containing glucose is subsequently fermented to ethanol using yeast. The second-generation bioethanol is produced from lignocelluloses, which are mainly composed of cellulose, hemicelluloses, and lignin. Cellulose and hemicelluloses are enzymatically hydrolyzed to glucose and pentoses and subsequently fermented to bioethanol (Naik et al., 2010). However, the complex structure formed by the polysaccharides and lignin prevents the accessibility of the polysaccharides to enzymatic attack (Menon & Rao, 2012). Therefore, the process involves a pretreatment of lignocelluloses to break the structural barrier. The pretreated lignocelluloses are enzymatically hydrolyzed to simple sugars using a cocktail of cellulases and hemicellulases. The simple sugars are subsequently fermented to ethanol using suitable ethanologens. The third-generation bioethanol is produced from algal biomass. The composition of algal biomass varies with species and the major polysaccharides present in algal biomass are glucans and galactans (Daroch et al., 2013). The algal feedstocks are chemically or enzymatically hydrolyzed to simple sugars, which are further fermented to ethanol. The enzymatic hydrolysis of algal biomass requires a pretreatment to improve the enzymatic hydrolysis (Lee, Kim, Um, & Oh, 2013; Nguyen,

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Choi, Lee, Lee, & Sim, 2009). Due to variation in the composition of algal feedstocks, enzymatic hydrolysis uses different polysaccharide-degrading enzymes depending on the composition of polysaccharides present in the algal species.

3. MARINE ENZYMES USED FOR BIOETHANOL PRODUCTION 3.1 Production and Biochemical Properties The enzymes used for bioethanol production vary with the feedstocks. The major enzymes used in the production of three generations of bioethanol are given in Table 1. Several marine microorganisms produce all these enzymes. The production of amylases by marine microbes has been reported from different genera including Streptomyces, Saccharopolyspora, Aureobasidium, Vibrio, and Pseudoalteromonas (Chakraborty et al., 2011; Chakraborty, Raut, Khopade, Mahadik, & Kokare, 2012; Li, Chi, Wang, & Ma, 2007; Najafi & Kembhavi, 2005; Tao, Jang, Kim, Yu, & Lee, 2008). The marine amylases exhibit tolerance to salts, alkali, and other reagents. Saccharopolyspora sp. amylase is tolerant to surfactants, oxidants, detergents, and salt (11% NaCl) (Chakraborty et al., 2011). The activity and stability of α-amylases produced by marine sponge are independent of calcium ions (Mohapatra, Banerjee, & Bapuji, 1998). This is contrast to other mesophilic terrestrial amylases, which are generally dependent on calcium ion for activity and stability. The α-amylase from marine bacterium Vibrio sp. is inhibited by EDTA and EGTA, while 25–55% of maximal activity is restored by divalent metal ions (Najafi & Kembhavi, 2005). The glucoamylase of marine yeast A. pullulans has a high molecular weight of 98 kDa with two subunits of 65 and 33 kDa. The glucoamylase is activated by most divalent ions (Li et al., 2007). The α-amylase (AmyZ) of Zunongwangia profunda is a cold-active enzyme with maximum activity at 35°C and retains about 39% activity at 0°C (Qin, Huang, & Liu, 2014). The AmyZ is salt tolerant with high activity at 1.5 M NaCl and 93% activity at higher concentration of NaCl (4 M). Cellulases are produced by several marine bacteria and fungi belonging to diverse genera including Bacillus, Dendryphiella, Saccharophagus, and Thermotoga maritima (Goyal, Selvakumar, & Hayashi, 2001; Kim et al., 2009; MacDonald & Speedie, 1982; Pointing, Vrijmoed, & Jones, 1998; Taylor et al., 2006). A marine protista Schizochytrium aggregatum has been reported to produce cellulase (Bremer & Talbot, 1995). Marine cellulases have optimal activity in a wide range of pH. The cellulase of S. aggregatum has activity at acidic pH 4.0–5.0, whereas the cellulase of

Table 1 The Major Enzymes and Some of the Feedstocks Used for the Production of Three Generations of Bioethanol First Generation Second Generation Third Generation Feedstocks

Enzymes

Feedstocks

Enzymes

Feedstocks

Enzymes

Wheat Corn Barley Rye Sorghum Potatoes Cassava Sweet potatoes

α-Amylase Pullulanases Glucoamylases Amyloglucosidase

Rice straw Wheat straw Sugarcane bagasse Corn stalk Saw dust Cotton stalk Elephant grass Napier grass Switch grass

Endo-β-1,4-glucanases Cellobiohydrolases Xylanases β-Glucosidases β-Xylosidases

Algae

Cellulase Carrageenase Agarase

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B. aquimaris is optimally active at alkaline pH 11 (Trivedi et al., 2011a). These cellulases exhibit optimal activity at mild temperatures like 30–40°C. Similarly, a cellulase from marine B. subtilis strain showed also high activity at low temperatures ranging from 20 to 40°C (Kim et al., 2009). While the cellulases are active in the presence of most common divalent metal ions, the activity is inhibited by Hg2+ like the terrestrial cellulases. The cellulase of marine bacterium Bacillus flexus is active at alkaline pH 9–12 and in the presence of 15% NaCl (Trivedi et al., 2011b). Similar to marine fungal and bacterial cellulases, a carboxymethyl cellulase of marine yeast A. pullulans is also optimally active at temperature of 40°C and the cellulase is active in the presence of Na+ and divalent metal ions (Yanjun, Liang, Zhenming, & Xianghong, 2015). However, the optimal pH and temperature for activities of cellulases from marine fungus D. arenaria are similar to most terrestrial fungal cellulases (MacDonald & Speedie, 1982). Xylanases are produced by several marine bacteria and fungi belonging to various genera such as Halorhabdus, Penicillium, Bacillus, Thermoanaerobacterium, Glaciecola, and Streptomyces (Guo, Chen, Sun, Zhou, & Zhang, 2009; Hou, Wang, Long, & Zhu, 2006; Hung et al., 2011; Liu, Zhao, & Bai, 2013; Menon, Mody, Keshri, & Jha, 2010; Waino & Ingvorsen, 2003). Though marine xylanases are similar to mesophilic terrestrial xylanases, some of them have specific properties like absence of cellulase activity, cold adaptation, and haloalkalitolerance. The cold-active xylanases have high activity at low temperatures (2–30°C) and are highly sensitive to high temperature (>50°C) (Marx, Collins, Amico, Feller, & Gerday, 2006). The xylanase of Glaciecola mesophila is a cold-active enzyme, which has an optimum temperature for activity at 30°C and retains 23% of optimal activity at 4°C. It has low thermostability retaining 20% of the activity after incubation for 60 min at 30°C (Guo et al., 2009). The marine haloalkalitolerant xylanases are also active in the presence of divalent cations such as Fe2+, Ca2+, and Mg2+, and the activity is greatly enhanced by these ions. The salt tolerance of marine xylanases varied widely. The xylanase produced from S. viridochromogenes is stable at NaCl concentration up to 5 M. Xylanase of T. saccharolyticum is highly active at NaCl concentration up to 12.5%. This enzyme is partially stable at high salt concentrations with retention of 67% of its activity after 48-h incubation in 15% (w/v) NaCl (Guo et al., 2009; Hung et al., 2011). The xylanases of B. pumilus and Halorhabdus utahensis are highly active at NaCl concentration up to 15% (Waino & Ingvorsen, 2003). Like other marine xylanases, these xylanases also retain 55% of the initial activity after 24-h incubation in high salt content like 20% NaCl.

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Carrageenase is produced by marine microbes of different genera including Vibrio, Cytophaga, Pseudomonas, Pseudoalteromonas, and Alteromonas (Araki, Higashimoto, & Morishita, 1999; Barbeyron, Gerard, Potin, Henrissat, & Loareg, 1998; Barbeyron, Henrissat, & Kloareg, 1994; Ohta & Hatada, 2006; Shangyong, Panpan, Linna, Wengong, & Feng, 2013). P. carrageenovora is the most studied bacterium for degradation of carrageenans. κ-Carrageenases have low molecular weight and are highly active around neutral pH and 45°C. Like other marine hydrolytic enzymes, the marine carrageenases are highly active in the presence of salts like NaCl and KCl. But, in contrast to marine cellulases, the activity of carrageenases is severely inhibited by divalent metal ions (Shangyong et al., 2013). Similar to cold acitve marine cellulases and xylanases, the κ-carrageenases are also active at mild temperatures. Carrageenase of Vibrio sp. is optimally active at 40°C and its activity is completely inhibited by the metal ions Zn2+, Pb2+, and Hg2+ and enhanced by Na+ (Araki et al., 1999). Several marine microbes of diverse genera including Pseudomonas, Pseudoalteromonas, Streptomyces, Vibrio, Alterococcus, Thalassomonas, and Saccharophagus are known to produce agarases (Ekborg et al., 2005; Kong, Hwang, Kim, Bae, & Kim, 1997; Lu et al., 2009; Meryandini, Junior, & Rusmana, 2011; Oh et al., 2010; Schroeder, Jaffer, & Coyne, 2003; Temuujin, Chi, Chang, & Hong, 2015). The marine agarases are also active at 30–40°C (Tao et al., 2008). An exo-β-agarase of an endophytic Pseudomonas sp. isolated from the red alga Gracilaria dura has a molecular mass of 66 kDa and exhibited optimal activity at 35°C. Its activity is stable at alkaline pH 7–11 and at high salt concentration up to 4 M (Gupta, Trivedi, Kumar, Reddy, & Jha, 2013).

3.2 Cloning and Characterization The genes encoding different saccharolytic enzymes of marine bacteria and fungi have been cloned and characterized. An amylase gene (amyA) encoding α-amylase from a marine bacterium Pseudoalteromonas sp. has an ORF of 2.007 kb encoding a polypeptide of 669 amino acids with a predicted molecular weight of 73.77 kDa and a pI of 5.15 (Tao et al., 2008). A novel gene (amyZ) encoding a cold-active and salt-tolerant amylase (AmyZ) of Z. profunda has a length of 1785 bp and encodes 66 kDa amylase of 594 amino acids (Qin et al., 2014). The AmyZ belongs to GH family 13 and is similar to α-amylase of Thermoactinomyces vulgaris. The amino acid sequence of the amylase showed 86% similarity to the α-amylase of Pseudoalteromonas haloplanktis.

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Cellulolytic genes have been cloned from many marine microbes and characterized. A 1.362 kb gene encoding a novel endo-β-1,4-glucanase (Cel5A) of Vibrio sp. has a catalytic domain of GH5 family and a cellulose-binding domain (CBM2). A β-glucosidase gene (bglB) of an extremely thermophilic eubacterium, T. maritima, has an ORF of 581 bp encoding 81 kDa protein (Gao, Ruan, Chen, Zhang, & Xu, 2010; Goyal et al., 2001). A putative carboxymethyl cellulase gene has been cloned from the marine yeast A. pullulans, which has a 954-bp ORF (Yanjun et al., 2015). A xylanase gene (xyn40) of a marine B. subtilis cho40 has an ORF of 645 bp encoding 215 amino acids with a molecular mass of 22.9 kDa. The sequence is homologous to that of B. subtilis xylanase with all features of xylanase enzyme. There are some differences with more serine residues and the presence of higher level of polar amino acids than nonpolar amino acids. The xynA gene of marine bacterium G. mesophila is 46% identical to the xylanase from Flavobacterium sp. strain MSY2. The gene (xynFCB) of xylanase from T. saccharolyticum NTOU1 belongs to GH10 family (Hung et al., 2011). Marine bacterium Saccharophagus degradans strain 2-40 has a versatile metabolism to degrade multiple polysaccharides present in different feedstocks of bioethanol (Taylor et al., 2006). Its genome has been fully sequenced and characterized. More than 180 ORFs encoding hydrolases degrading polysaccharides are identified in the genome. Specifically, the genome has a complete set of genes encoding cellulolytic and hemicellulolytic enzymes for degradation of plant biomass. A κ-carrageenase gene (cgkZ) of marine bacterium Zobellia sp. ZM-2 has an ORF of 1638 bp encoding 545 amino acids with 35 amino acids signal peptide (Liu et al., 2013). The structural gene cgkA encoding κ-carrageenase Alteromonas carrageenovora codes for 397 amino acids, with a signal peptide of 25 amino acids (Barbeyron et al., 1994). The carrageenase CgkA belongs to a new member of family GH16, which comprises β-1,3-1,4-glucanases from various sources and the β-agarase of Streptomyces coelicolor (Temuujin et al., 2015). The residues Glu163 in the κ-carrageenase CgkA and Glu155 in the β-agarase from S. coelicolor are proposed to play an important role in catalysis (Barbeyron et al., 1994). A κ-carrageenase gene of marine gliding bacterium Cytophaga droebachiensis consists of an octameric omega region similar to the ribosomal-binding site (Barbeyron et al., 1998). The agarase gene agaA of marine bacterium Pseudomonas vesicularis MA103 has 2958 bp coding for 985 amino acids with a predicted signal peptide of 29 amino acids. The AgaA has a GH16 catalytic domain and three carbohydrate-binding modules 6 (Kong et al., 1997). The agarase gene (agaA6) of Pseudoalteromonas sp. JT-6

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has an ORF of 1338 bp encoding 445 amino acids with a predicted Mr of 50 kDa (Tao et al., 2008). The amino acid sequence of AgaA6 consisting of a signal peptide, a catalytic domain of family GH16, and a carbohydrate-binding domain is 99% identical to the AgaA of Janthino sp. SY12 (Tao et al., 2008). The β-agarase of the marine Vibrio sp. also belongs to GH16 family (Zhang & Sun, 2007). A 0.87 kb β-agarase gene (agrP) of Pseudoalteromonas sp. AG4 encodes 290 amino acids with a signal peptide of 21 amino acids, which is 98.6% identical to β-agarase of P. atlantica (Tao et al., 2008). The β-agarase gene agaA of Pseudoalteromonas sp. CY24 has an ORF of 1359 bp encoding 453 amino acids comprising a catalytic domain of family GH16 and a carbohydrate-binding module type 13 (Lu et al., 2009). The residues V109VTS112 play a key role in the enzyme catalysis.

4. MARINE MICROOORGANISMS PRODUCING ETHANOL Marine yeasts tolerant to salt are known to produce ethanol. These yeasts are useful for the fermentation of high salt sugar medium. Marine yeasts such as Candida sp. and Debaryomyces sp. isolated from coastal water have higher salt tolerance (2.0–3.0 M NaCl) compared to that of industrial yeasts (Urano, Hirai, Ishida, & Kimura, 1998). A marine Saccharomyces cerevisiae is capable of producing ethanol at a yield of 69.58%. Protoplast fusion has been followed to generate efficient halotolerant ethanologenic yeasts. A high ethanol producing flocculent S. cerevisiae on hybridization with a halotolerant nonflocculent Zygosaccharomyces rouxii by polyethylene glycol-induced protoplast fusion has generated fusants with ability to produce ethanol in salted medium (Limtong, Deejing, & Yongmanitchai, 1998). The fusants are able to produce 6.85% ethanol with 87% theoretical yield from glucose broth containing NaCl up to 7%. The halotolerant ethanologenic yeasts play an important role in fermentation of the hydrolysate of marine biomass, since the marine feedstocks result in a salty hydrolysate, which prevents the use of conventional ethanologens (Khambhaty et al., 2012). The use of marine halotolerant ethanologens for direct fermentation of algal hydrolysates can avoid the use of the energy intensive desalting step, which can lead to economic process. In addition, the halotolerant yeasts can be used for the production of bioethanol of first and second generations by involving seawater in the process. The use of seawater prevents the exploitation of freshwater in bioethanol industry. It has been estimated that the production of bioethanol by first- and second-generation technology

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consumes 2.7–5.8 gallons of freshwater per gallon of bioethanol produced (Wu & Chiu, 2011). The large volume of freshwater can be saved by using seawater as an alternative water resource for the production of bioethanol in coastal areas. Therefore, the seawater-based process can reduce depletion of freshwater reservoir. The added advantage is that the minerals present in the seawater can support the growth of ethanologens during fermentation (Lin, Luque, Clark, Webb, & Du, 2011). This can further minimize the addition of nutrients to the fermentation medium. Hence, the use of seawater in bioethanol process has the potential to improve the economics of the process with minimal environmental emissions. The marine halotolerant ethanologenic microorganisms play a very significant role for successful achievement of seawater-based bioethanol production process.

5. BIOETHANOL PRODUCTION FROM THE MARINE ALGAE Marine algae are commonly known as seaweeds. The marine algae include red, brown, and green algae (Ge et al., 2011). The marine algae are considered as an alternative renewable source for bioethanol production (Lee & Lee, 2016). Production of ethanol by utilizing green, red, and brown algal feedstocks has been reported. Various algal species belonging to these three types used for bioethanol production are shown in Table 2. There are several advantages of using marine algae for renewable bioethanol production (Fasahati & Liu, 2016). Algae have higher photon conversion efficiency and can accumulate high levels of polysaccharides (Daroch et al., 2013). The carbon dioxide tolerance and its rate of utilization are very high, which enables the algae to utilize carbon dioxide emitted from fossil fuel-based power stations and chemical industries (Cuellar-Bermudez, Garcia-Perez, Rittmann, & Parra-Saldivar, 2015). The consumption of carbon dioxide emitted from industry would further significantly reduce green house gas level in the atmosphere. There are a very large number of species of algae with varying properties, which enables selection of species suitable to the regional setup and the environment. The feedstock supply and sustainability for bioethanol production are relatively higher for algal biomass, since algae have the capacity to produce high quantity of biomass compared to plants and the biomass production time is short for algae; hence, multiple harvests are possible annually compared to plants (Ge et al., 2011). This would avoid the scarcity of biomass and fulfill the required production capacity, which can allow continuous supply of bioethanol as per the demand.

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Table 2 Some Marine Algal Feedstocks and Methods of Conversion to Bioethanol Algal Feedstocks Pretreatment Ethanologen Yield (%) References

Chlorococcum infusionum

S. cerevisiae

26.00

Harun, Jason, Cherrington, and Danquah (2011)

Chlamydomonas Hydrothermal acid, reinhardtii α-amylase (90°C, 30 min), glucoamylase UTEX 90 (55°C, 30 min)

S. cerevisiae S288C

23.50

Nguyen et al. (2009)

Kappaphycus alvarezii

0.9 N sulfuric acid, 120°C, 60 min

S. cerevisiae 15.40 NCIM 3455

Sargassum sagamianum

Thermal liquefaction, P. stipitis 200°C, 15 MPa, 15 min CBS7126

Laminaria japonica

0.1 N HCl, 121°C, E. coli KO11 16.10 15 min, Celluclast 1.5 L

Lee et al. (2013)

Gracilaria salicornia

2% sulfuric acid, 120°C, E. coli KO11 7.90 30 min, cellulase

Wang, Liu, and Wang (2011)

Chlorella vulgaris

0.1–5.0% (v/v), sulfuric Z. mobilis acid, 121°C, 20 min

Ho et al. (2013)

0.75% (w/v) NaOH, 120°C, 30 min

10.00

1.12

Khambhaty et al. (2012) Yeon et al. (2011)

The major technical advantage of seaweeds is the lack of lignin in the biomass, which makes the biomass hydrolysis easier compared to lignocellulose hydrolysis (Ge et al., 2011). However, unlike lignocellulose, the type of polysaccharides present in the seaweeds varies with the species. There are several species each with different sugar compositions. The composition of acid and enzyme hydrolyzate of different marine algae, Ulva lactuca, Gelidium amansii, Laminaria japonica, and Sargassum fulvellum, contained multiple sugars such as glucose, mannose, galactose, and mannitol at different ratios. Fermentation of multiple sugars in the hydrolysate of algal biomass has been studied using yeasts (Fasahati, Woo, & Liu, 2015; Khambhaty et al., 2012; Yanagisawa, Nakamura, Ariga, & Nakasaki, 2011; Yoza & Masutani, 2013). A high yield of bioethanol production from Ulva fasciata has been reported, where the biomass is enzymatically hydrolyzed using cellulase from Trichoderma reesei. The ethanol production has been reported at 0.45 g/g sugar with 88.2% theoretical yield and 90 L/t of dry biomass (Trivedi, Gupta, Reddy, & Jha, 2013). Ethanol production from brown algae L. japonica has been reported at a concentration of 2.59 g/L

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(Lee et al., 2013). Fermentation of multiple sugars in L. japonica hydrolysate supplemented with Luria-Bertani medium by ethanogenic recombinant Escherichia coli KO11 produced 0.4 g ethanol per gram of carbohydrate (Kim, Li, Jung, Chang, & Lee, 2011). Ethanol production from algal biomass Saccharina japonica has been studied using a recombinant E. coli, transformed with a cluster of genes and transporter involved in alginate, mannitol, and glucose metabolism, through consolidated bioprocessing (Wargacki et al., 2012). This method shows the utilization of alginate, mannitol, and glucan with ethanol production at a yield of 0.41 g/g sugars. Another ehtanologenic yeast Brettanomyces custersii, capable of utilizing glucose and galactose, has been used to produce bioethanol from red seaweeds G. amansii (Yeon et al., 2011). There are few reports on bioethanol production from brown seaweeds by simultaneous saccharification and fermentation. The simultaneous saccharification and fermentation of Saccharina produced ethanol at 7.7 g/L (Jang, Cho, Jeong, & Kim, 2012). The fermentation of alginate from L. japonica by ethanologens isolated from nuruk showed utilization of alginate (Lee & Lee, 2011). The utilization of alginate and mannitol by the industrial ethanologen S. cerevisiae has been achieved by genetic engineering (Enquist-Newman et al., 2014). The engineered yeast produced ethanol from mannitol and alginate with a theoretical yield of 83%. There is a need for further development of engineered microbes for efficient utilization of algal biomass for bioethanol production.

6. CONCLUSION Production of fuel ethanol has attracted attention worldwide due to the oil crisis that began a few decades ago. Considering the sustainability, economic, and environmental advantages of fuel ethanol, the global production of ethanol has increased and it has been supported by policies and incentives. But the growth of fuel-ethanol production is hindered by hurdles related to different factors including conversion technology. One of the major factors is unavailability of efficient enzyme systems for the process based on low energy and emission. Current enzymes are optimized to work optimally at temperatures of 50°C, but the hydrolysis conditions are not compatible with optimal fermentation conditions, where lower temperature (30–37°C) is used. Some of the main issues in bioethanol production could be addressed by employing marine biocatalysts. The hydrolysis and fermentation efficiency with respect to product yield and energy usage could be improved by using marine cold-adaptive enzymes. Another major concern

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is freshwater usage because its consumption by bioethanol industries is projected to increase significantly by 2050. This would have a severe impact on agriculture and livestock that could be avoided by exploiting the potentials of halotolerant marine biocatalysts as an alternative to the conventional biocatalysts in the bioethanol process. Application of metabolic engineering to tune the physiology of marine organisms would result in achieving the goal of bioethanol production. Though cellulosic bioethanol is renewable, the production of the second-generation bioethanol is costly because of the high recalcitrance of lignocellulosic raw materials. Whereas marine algae are not recalcitrant like lignocelluloses. Therefore, the algal biomass as feedstocks for bioethanol production is a promising source for less-energy-intensive renewable bioethanol production. Eventually, the cultivation of algae in ocean would leave the agricultural land for cultivation of food crops and avoid the fuel–food conflict.

ACKNOWLEDGMENTS We acknowledge IIT Madras for awarding Post Doctoral Fellowship and Half Time Teaching Assistantship to Dr. Manas and Vignesh, respectively.

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CHAPTER TEN

Enzymes in Fermented Fish Giyatmi*, H.E. Irianto†,1 *Jakarta Sahid University, Jakarta Selatan, DKI Jakarta Province, Indonesia † Center for Fisheries Research and Development, Jakarta Utara, DKI Jakarta Province, Indonesia 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Enzymes in Fish 3. Enzymes in Fermented Fish 3.1 Endogenous Fish Enzymes 3.2 Microbial Enzymes 3.3 Enzymes Added to the Fermentation Process 4. Enzymes in Fish Sauce Processing 5. Concluding Remarks References

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Abstract Fermented fish products are very popular particularly in Southeast Asian countries. These products have unique characteristics, especially in terms of aroma, flavor, and texture developing during fermentation process. Proteolytic enzymes have a main role in hydrolyzing protein into simpler compounds. Fermentation process of fish relies both on naturally occurring enzymes (in the muscle or the intestinal tract) as well as bacteria. Fermented fish products processed using the whole fish show a different characteristic compared to those prepared from headed and gutted fish. Endogenous enzymes like trypsin, chymotrypsin, elastase, and aminopeptidase are the most involved in the fermentation process. Muscle tissue enzymes like cathepsins, peptidases, transaminases, amidases, amino acid decarboxylases, glutamic dehydrogenases, and related enzymes may also play a role in fish fermentation. Due to the decreased bacterial number during fermentation, contribution of microbial enzymes to proteolysis may be expected prior to salting of fish. Commercial enzymes are supplemented during processing for specific purposes, such as quality improvement and process acceleration. In the case of fish sauce, efforts to accelerate fermentation process and to improve product quality have been studied by addition of enzymes such as papain, bromelain, trypsin, pepsin, and chymotrypsin.

Advances in Food and Nutrition Research, Volume 80 ISSN 1043-4526 http://dx.doi.org/10.1016/bs.afnr.2016.10.004

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1. INTRODUCTION Fermented fish products are very popular in Southeast Asian countries; however, the products are actually also found in other parts of the world. Fermentation of fish is an ancient technology that has already been employed by our ancestors a long time ago. The processing is traditionally used to overcome the perishable nature of fish. Fermented fish is an old staple food in European cuisines; for instance, the ancient Greeks and Romans made a famous fermented fish product called “garum.” The product has pasta form and very strong smell. Garum is made through a fermentation process of entrails and blood of mackerel (Ching, Mauguin, & Mescle, 1992; Gildberg, Simpson, & Haard, 2000; Ska˚ra, Lars Axelsson, Stefansson, & Ekstran, 2015). Fermented fish products usually have special consumers because of their ability to provide a certain unique characteristic, especially in terms of aroma, flavor, and texture. This is due to a transformation of organic materials into compounds which are simpler by the activity of microorganisms or enzymes that are encountered in fish muscle tissue during the fermentation process (Beddows, 1998). The interest of consumers for fermented fish products is primarily due to the specific flavor generated which can induce appetite. In the case of Indonesia, a variety of flavors produced by fermented fish products can actually satisfy the tastes of consumers outside the area of origin of the product. Unfortunately most of fermented fish products are still local and not so easily found nationwide. Only some types of fermented fish products have been widely known, such as fish sauce and shrimp paste (Irianto, 2012). The ones contributing the most in flavor formation and the changes in texture in fermented fish products are enzymes. Besides enzymes, microorganisms that contribute to the fermentation process also assist in the formation of aroma and flavor (Beddows, 1998). Many researchers from around the world have already explored the enzymes in fermented fish products and pay attention to uncover their role in the fermentation process.

2. ENZYMES IN FISH Naturally, fish contains enzymes which are distributed in the whole body of the fish. The blood, certain tissues, muscles, and glands, such as kidney, contain very active enzymes (Marsh & Flick, 2012). Enzyme action

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often causes significant deteriorative changes prior to noticeable spoilage of bacterial origin (Ghaly, Dave, Budge, & Brooks, 2010). Autolytic enzymes are present at much higher concentration in the viscera and head than in other tissues (Owens & Mendoza, 1985). During rigor mortis, the acidity of the tissues decidedly increases and this change of the hydrogen ion concentration causes an acceleration of autolytic decomposition. Enzymes are most active in dilute solution and do not act in the absence of water. Most of them are destroyed or rendered inactive by concentrated salt solution (Gildberg et al., 2000; Ponce de Leon Valdivia, 1994). The proteolytic activity is mainly caused by tissue proteases and, to a lesser degree, by gut proteases (Lindgren & Pleje, 1983). Based on the mode of catalysis, proteolytic enzymes from the fish may be classified into: (a) aspartic proteases (pepsin and cathepsin D); (b) serine proteases (trypsin and chymotrypsin); (c) cysteine proteases (calpain, m-calpain, and cathepsins B, H, L); and (d) metalloprotease (Sriket, 2014). Aspartic and alkaline proteases have been found from the viscera and stomach of sardine (Bougatef, Souissi, Fakhfakh, Ellouz-Triki, & Nasri, 2007; Khaled et al., 2011; SalazarLeyva et al., 2013). Klomklao (2008) proposed proteases to be classified on the basis of their similarity to well-characterized proteases, such as trypsin-like, chymotrypsin-like, chymosin-like, or cathepsin-like. Trypsins play major roles in biological processes including digestion, activation of zymogens of chymotrypsin, and other enzymes (Cao, Osatomi, Hara, & Ishihara, 2000). Serine collagenases or trypsin-like proteinase were found in the intestines of Atlantic cod, Gadus morhua (Hernandez-Herrero, Duflos, Malle, & Bouquelet, 2003), and king crab, Paralithodes camtschaticus (Rudenskaya, Kislitsin, & Rebrikov, 2004). Chymotrypsin was isolated from the hepatopancreas of Chinese shrimp, Fenneropenaeus chinensis (Shi, Zhao, & Wang, 2008). Chymosin has been isolated and identified from carp and harp seal stomach (Shahidi & Kamil, 2001). Cathepsins also known as lysosomal cysteine proteases, playing an important role in many physiological processes including protein degradation, are mostly active at weakly acid pH values (pH 5). Cathepsins B, C, H, L, and S have been extracted from fish and shellfish muscles to be the major proteases involved in intracellular protein breakdown (Aoki, Ahsan, & Watabe, 2004; Pangkey et al., 2000). Cathepsin D shows some activity in the lowest pH range, predominating postmortem in some fish. The activity of cathepsin D was detected in red or white fish muscle among 24 species, and no difference was found between red- and white-flesh fish, or freshwater fish (Aoki, Yamashita, & Ueno, 2000).

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A study on digestive enzymes conducted by Langeland, Lindberg, and Lundh (2013) proved that the high lipase and protease activity and low carbohydrase activity in Eurasian perch (Perca fluviatilis) and Arctic charr (Salvelinus alpinus) can be linked to their feeding habits. Total carbohydrase activity was higher in Eurasian perch than in Arctic charr, which had a higher total chymotrypsin activity and lipase activity in the mid-intestine. The study suggests that feed formulation should be different for Eurasian perch and Arctic charr in order to match their inherent digestive enzyme activities. This relates specifically to the carbohydrate content and composition of the feed. Based on the observed α-amylase and carbohydrase activities, the carbohydrate content, particularly starch, can be higher in feed for Eurasian perch than in feed for Arctic charr. In fish, the levels of digestive enzymes may be influenced by type of feeding (Hofer & Schiemer, 1981), the age of the fish (Il’ina & Turetskiy, 1988), season and/or temperature of acclimatization (Kuz’mina, 1991), and so on. Additionally, heavily feeding fish will generally deteriorate rapidly because the enzyme concentration is often higher in the digestive tract of the fish during feeding. The effect of season on enzyme activity varies with the spawning cycle, water temperature, feeding cycle, and other variables (Wheaton & Lawson, 1985). The activity and thermal stability of fish enzymes vary from one species to another. For example, the activity and thermal stability of tryptic enzymes from horse mackerel (Trachurus mediterraneus ponticus) are greater than those from sprat (Sprattus nostamus). The pepsin from plaice (Pleuronectes platessa) is 10 times more active than that from horse mackerel. Generally, white fish have less proteolytic activity than do pelagic species (Mackie, Hardy, & Hobbs, 1971). The presence of active proteases in muscle and digestive organ, particularly proteolytic and collagenolytic enzymes, makes the flesh fish and shellfish prone to degrade especially during storage in ice, since the digestive organ is not practically removed prior to storage. That occurrence leads to the muscle softening or mushiness of fish and shellfish during storage or distribution. During storage of fish and shellfish, the intensive hydrolysis of myofibrillar and collagenous proteins by proteases was noted. To lower the muscle degradation, different pretreatment methods as well as protease inhibitors have been applied in the stored fish and shellfish. The use of natural serine or trypsin inhibitors is a better way to retard such a textural problem of fish species (Sriket, 2014). Biochemical properties of trypsin purified from the digestive system of carp Catla catla had a similarity with trypsin from other fishes. Stability at high pH and low temperature indicates the potential application of this

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protease in detergent and in the food industry. Enzymes extracted from the fish viscera may be used in the food processing industry, thus making beneficial and productive use of the fish processing wastes (Khangembam, Sharma, & Chakrabarti, 2012). Each type of fish protease extracted from the visceral waste of different fish species had a distinct optimum alkaline pH, temperature, and molecular weight. The crude enzyme from the visceral waste of the Red snapper and Great barracuda can be used to remove the blood stains effectively within 20 min, without the usage of any detergents. It was also observed that the enzyme dehaired the goat hide after 22 h of incubation, without the addition of sodium sulfide. The direct applicability of the crude extract without downstream processing would make its use acceptable as a substitute for the commercial ones (Sabtecha, Jayapriya, & Tamilselvi, 2014). Atlantic cod as a poikilothermal organism lives in relatively frigid water; thus, it can be predicted that its enzymes are classified as cold active enzymes. Several of these enzymes have been demonstrated to be cold active, including elastase, collagenase, and chymotrypsin. The enzyme activity at low temperatures is helpful in various food processing applications where proteolysis must be performed at low temperature, such as in caviar production and carotenoprotein extraction (Vilhelmsson, 1991). Fish also contains transglutaminase, playing an important function in surimi production. The role of transglutaminase in surimi production is in the formation of ε-(γ-glutamyl) lysine bonds in the fish protein to produce a high-quality gel. The activity of endogenous fish transglutaminase decreases rapidly after catch, and is almost completely destroyed by freezing (Vilhelmsson, 1991).

3. ENZYMES IN FERMENTED FISH Proteases play an important role in the production of fermented fish product, particularly during fermentation process to obtain an acceptable product quality. Chadong, Yunchalard, and Piyatheerawong (2015) demonstrated fish protein degradation and peptide formation throughout fermentation process of Plaa-som, a traditional fermented fish product from Thailand, due to proteolysis. Proteolysis of Plaa-som occurred under fermentation at 30°C because protein concentration was continually reduced to 8.52 mg/g after 120 h of fermentation. This fact was also supported by the increase of peptide content to 1.95 μmol/g during fermentation.

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Fermentation process of fish relies both on naturally occurring enzymes (in the muscle or the intestinal tract) as well as those enzymes from bacteria. In general, the former is most considerable with respect to changing texture as well as producing some of the flavor, and the latter contributes to the development of aroma and flavor (Ska˚ra et al., 2015). In some cases, commercial enzymes are also supplemented for specific purposes, such as quality improvement and process acceleration.

3.1 Endogenous Fish Enzymes As discussed above, various proteolytic enzymes are found in viscera, digestive tract, and muscle tissue of fish. Major endogenous proteinases in anchovies were trypsin-like proteinase, pepsin, chymotrypsin, elastase, and aminopeptidase (Martinez & Serra, 1989; Siringan, Raksakulthai, & Yongsawatdigul, 2006). Digestive enzymes of trypsin, chymotrypsin, and pepsin are considered as the three of more important enzymes compared to others (de la Parra, Rosas, Lazo, & Viana, 2007). Pepsin is usually found in the stomach of fish and is a main enzyme of the digestive juices (de la Parra et al., 2007). Trypsin is present in viscera, pyloric caeca, and spleen (Kishimura, Hayashi, Miyashita, & Nonami, 2005, 2006; Kishimura et al., 2007; Klomklao et al., 2006). Hepatopancreas of fish and shellfish digestive organs contains both peptidase and proteinase activities such as aminopeptidase, gelatinolytic proteases, trypsin and chymotrypsin, and collagenolytic proteases (Sriket, 2014). It was found that most of lipolysis and proteolysis activities in peda processing were recorded in the gut, especially at the beginning of the fermentation process; however, the activities fell rapidly during the process (Irianto, 1990). In view that enzymes are found in viscera and digestive tract, evisceration plays an important role in determining the rate and type of enzymatic degradation occurring. Fermented fish products processed using the whole fish will have different characteristic than those manufactured from headed and gutted fish (Wheaton & Lawson, 1985). The enzymatic activity of most visceral and digestive tract enzymes from fish had the greatest activity at near neutral pH values (Bougatef et al., 2007; Munilla-Moran & Saborido-Rey, 1996). Castillo-Yan˜ez, Pacheco-Aguilar, Garcia-Carren˜o, and Toro (2004) isolated an acid proteolytic enzyme, which belongs to the aspartic protease class from the viscera of sardines. The enzyme is similar to pepsin II from other fish species and is stable at pH 3–6 and 45°C.

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The pyloric caeca represent the organs which are the major source of alkaline proteinases. A trypsin-like enzyme obtained from the pyloric caeca of cod (G. morhua) had an isoelectric point of 5.30 and 5.89 and was very similar in amino acid composition to bovine trypsin, but differed in having a higher relative amount of acidic amino acids and a lower amount of basic amino acids. The enzyme also hydrolyzed fish protein substrates (Beira˜o, Mackie, Teixeira, & Damian, 2001). Three alkaline proteinases and two acid proteinases were isolated from sardine. Each of the alkaline proteinases hydrolyzed casein more rapidly than other proteins. A major alkaline proteinase (III) hydrolyzed sarcoplasmic proteins from sardine five times faster than other alkaline proteinases. Each of two acid proteinases hydrolyzed hemoglobin and myoglobin more rapidly than the other proteins. After preincubation with 25% NaCl, an alkaline proteinase (III) and an acid proteinase (II) were stable although the other proteinases became unstable. The two proteinases, alkaline proteinase III and acid proteinase II, were also stable for 3 months after the beginning of fish sauce production. The proteolytic activity of each of alkaline and the acid proteinases was strongly inhibited by more than 15% NaCl; however, minimum inhibition was observed when sardine muscle proteins were used as substrate (Noda, Van, Kusakabe, & Murakami, 1982). Two aminopeptidases (I and II) were extracted from defatted internal organs of sardine and purified using DEAE-cellulose chromatography, gel filtration on Sephadex G-200, and isoelectric focusing. The final preparations of enzymes I and II were judged nearly homogenous by polyacrylamide gel electrophoresis. The molecular weights of enzymes I and II were determined by gel filtration to be 370,000 and 320,000, respectively. The isoelectric points were 4.1 (I) and 4.8 (II), respectively. Both enzymes were inhibited by EDTA and activated by Co++. Bestatin could inhibit enzyme I but not enzyme II. Enzymes I and II rapidly hydrolyzed not only synthetic substrates containing alanine or leucine but also di-, tri-, and tetraalanine. Based on all of these characteristics, sardine aminopeptidases resemble human alanine aminopeptidase. Enzyme I retained more than 70% of its original activity in 15% NaCl, suggesting the enzyme participates in hydrolyzing fish proteins and peptides during fish sauce production (Vo Van, Kusakabe, & Murakami, 1983). Activities of alkaline and acid proteinases were compared with bovine trypsin and pepsin and showed that like bovine trypsin the alkaline proteinase from sardines pyloric caeca hydrolyzed casein more effectively than other protein substrates (Noda et al., 1982).

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Muscle tissue enzymes, particularly cathepsins, peptidases, transaminases, amidases, amino acid decarboxylases, glutamic dehydrogenases, and related enzymes, are all found in fish muscle tissue (Chaveesuk, 1991), and these enzymes, particularly trypsin, chymotrypsin, and cathepsin, involve in the protein hydrolysis during fish sauce fermentation (Fernandes, 2016). Muscle tissue enzymes are mostly located in the cells. On the other hand, digestive enzymes are exocellular secretion. Even though some studies showed that muscle tissue enzymes have an optimum activity at neutral pH, most reports inform that low pH values accelerate muscle tissue enzyme activities. Most fermented fish products are processed at pH above 4, except for fish silage and some fermented fish products. Accordingly, most muscle tissue enzymes are actually not at optimum pH condition (Mackie et al., 1971). Partial characterization of cathepsins B from the muscle of horse mackerel indicated similar characteristics with other cathepsin BS. The optimum pH of the cathepsin was 5 with optimum temperature of 50°C. The activity was inhibited by E-64, CA-074, and chymostatin (Yoshida et al., 2015). Maximum enzyme activities can be achieved by using whole fish including heads and viscera. On the contrary, minimum enzyme activity will occur when deheaded and gutted fish are used to produce fermented fish products. Meanwhile, intermediate enzyme activities can be obtained by removing the guts anytime after the fish are caught to allow some diffusion of visceral enzymes into tissues (Owens & Mendoza, 1985). In salted fish, the ripening is described by three hypotheses. These are (1) microbiological theory, (2) autolytic theory, and (3) enzymes theory. In microbiological theory, the microorganisms produce the essential active enzymes, and these enzymes penetrate into the flesh and contribute to ripening process. The autolytic theory describes that ripening is a result of the activity of enzymes of the muscles or other tissues, or of the gastrointestinal tract. Finally, the enzyme theory explains the ripening of salted fish as taking place under the influence of certain enzymes, namely, those contained in the muscle tissue, those in the intestinal body organs of the fish, together with those produced by microorganisms (Mackie et al., 1971). In the maturing of anchovies, maximum autolytic activity of Indian anchovy (Stolephorus indicus) was found at 60°C. Autolytic activity decreased with increased NaCl concentration. Crude extract exhibited an optimum pH at 8.5–9.5. Trypsin-like proteinases were the predominant proteinases in the crude extract. Proteinases from Indian anchovy could participate in protein hydrolysis during fish sauce fermentation. Therefore, incubation of Indian anchovy at 60°C and in 10% NaCl for a period of time before full

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salting at 25% NaCl could be an effective way to accelerate the fish sauce fermentation process (Siringan et al., 2006).

3.2 Microbial Enzymes Fermentation of fish is brought about by autolytic enzymes from fish and microorganisms in the presence of salt. The use of salt in fresh fish preservation is as selective microbial agent (Anihouvi, Kindossi, & Hounhouigan, 2012; Majumdar & Basu, 2010). There are two categories of fermented fish products. They are (1) product preservation primarily by water activity reduction (fish/salt formulation) and (2) product preservation by combined water activity reduction and lactic acid generation (fish/salt/carbohydrate formulation) (Adam, Cooke, & Rattapol, 1985). During salting, genus Micrococcus is predominant and its proportion increases gradually from 40% to approximately 90% of the total numbers. Simultaneously an appreciable reduction is observed in the other genera which are named in the following order of importance: Flavobacterium, Achromobacter, Pseudomonas, Bacillus, and Sarcina (Graikoski, 1973). The salting treatment reduces the water activity of fish from about 0.98 to about 0.70–0.75. At this water activity range, the only possibility is the growth of halophilic bacteria, xerophilic molds, and osmophilic yeast (Grant, 2004; Smith, 1989; Stevenson et al., 2015). Microbial load in fish sauce made from gambusia (Affinis affinis) during processing decreased with increased fermentation period, possibly caused by high concentration of salt (Ibrahim, 2010). While similar phenomenon was noted, the growth of halophilic bacteria, lipolytic bacteria, proteolytic bacteria, and lactic acid bacteria in peda processing decreased during the first and second fermentation. The bacteria probably contributed significantly only at the beginning of the fermentation, since the number tended to decrease (Irianto, 1990). The role of microorganisms in the fermentation process of fish is different from that in fermented vegetable products. The high salt content of these products leaves only salt-tolerant microorganisms to survive (Rose, 1982). Microorganisms excrete proteolytic enzymes capable of degrading proteins. Many types of microorganisms excrete proteolytic enzymes, including the fungi Aspergillus oryzae, A. orizae, and Rhizopus sp.; the bacteria Bacillus subtilis; the actinomycetes Streptomyces griseus; and the yeast Saccharomyces spp. and Candida sp. (Gupta, Beg, Khan, & Chauhan, 2002; Mackie et al., 1971; Oyeleke, Egwim, Oyewole, & John, 2012). Therefore, careful selection by seeding or controlling the growth environment within the fermentation

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container enables the desired microbes to flourish and produce significant quantities of proteolytic enzymes which help to hydrolyze the fish protein (Wheaton & Lawson, 1985).

3.3 Enzymes Added to the Fermentation Process The fermentation process in the production of traditional fermented fish products is often found running slow, inconsistent product quality, and frequently not as expected. Microorganisms producing enzymes require a long adaptation period, particularly adjustment to environmental conditions with high salt content. However, the process acceleration, consistent product quality guarantee, and product quality improvement can be performed by using enzymes addition during processing. The enzymes may be produced from plant, animal, and microorganisms. Several enzymes are extracted from plant, and those have been well recognized for their ability to tenderize meats. Commercial plant proteases such as bioprase, pronase, molsin, protease AJ, papain, bromelain, and ficin have all been investigated as enzyme supplements to speed up the rate of fish sauce fermentation (Chaveesuk, 1991; Le et al., 2015; Ooshiro, Ok, Une, Hayashi, & Itakura, 1981; Yongsawatdigul, Rodtong, & Raksakulthai, 2007). Bromelain found in pineapple juice, papain from papaya latex, and ficin from figs are already well known (Wheaton & Lawson, 1985). Those enzymes are quite heat stable and work optimally at near neutral pH (Mackie et al., 1971). Proteases which have been investigated to be used in processing fermented fish products are extracted from A. oryzae (Man & Tuyet, 2006), moderately halophilic marine bacterium Pseudomonas sp. (Nakano, Watanabe, Hata, Qua, & Miura, 1986), and others. The most common commercial microbial proteases are Alcalase®, Neutrase®, Protamex®, Flavourzyme®, and Kojizyme® (Aristotelis, Himonides, Anthony, Taylor, & Morris, 2011; Nilsang, Lertsiri, Suphantharika, & Assavanig, 2005; Yongsawatdigul et al., 2007). Animal-derived enzymes are trypsin, pepsin, and pancreatin. Trypsin has been extracted from digestive system of carp C. catla (Khangembam et al., 2012). Fish pepsins have been purified and characterized in various types of fishes including Arctic capelin, rainbow trout, Atlantic cod, bolti fish, Antarctic rock cod, sea bream, African coelacanth, Mandarin fish, smooth hound, orange-spotted grouper, albacore tuna, and European eel (Zhao, Budge, Ghaly, Brooks, & Dave, 2011).

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4. ENZYMES IN FISH SAUCE PROCESSING A fish sauce has attracted research scientists of all over the world to explore the secrets behind its fermentation process. Fish sauce is not only familiar for the people in Southeast Asia but also for those living in other parts of the globe. Fish sauce is known as “kecap ikan” in Indonesia, “nam pla” in Thailand, “patis” in Philippines, “shottsuru” in Japan, “nu€ oc m^am” in Vietnam, “budu” in Malaysia, “ngapi” in Myanmar, “pissala” in France, “garos” in Greece, “colombo-cure” in Pakistan and India, “yeesu” in China, and “aekjeot” in Korea. Fish sauce is manufactured through fermentation process for 3–12 months, in which fish and salt are previously mixed thoroughly at a ratio of 1:3. After 4–6-month period, a liquid containing fish extract is obtained in fermentation tanks. That liquid is actually fish sauce. During fermentation process, fish tissue is gradually hydrolyzed, indicating the activity of proteolytic enzymes. The proteolytic enzymes responsible for the protein degradation are either endogenous fish enzymes coming from viscera or enzymes from microorganisms which may previously exist on or in the fish prior to the salting period. Endogenous proteolytic enzymes of fish originate from the digestive tract, internal organs, or muscle tissue (Chaveesuk, 1991; Chayovan, Rao, Liuzzo, & Khan, 1983). However, Orejana and Liston (1981) claimed that endogenous fish enzymes are the major and perhaps sole agents responsible for digestion in the fish sauce process. Fen, Sali, Ahmad, Tze, and Abdullah (2011) revealed similar finding that the endogenous fish enzymes, especially from fish viscera, were the main contributors of protease action during the initial days of fermentation. In addition, bacterial enzymes may be involved in the later stage of fermentation. Digestive enzymes have a significant role in the fermentation of capelin (Mallotus villosus) sauce, in view of the fact that the rate of protein hydrolysis of whole fish was considerably higher than that of eviscerated fish. Intracellular enzyme of cathepsin C was believed to contribute to proteolysis in fish sauce and the formation of the delicious fish sauce taste (Raksakulthai, 1987). Quality improvement and fermentation process acceleration of fish sauce can be carried out enzymatically through the use of papain (Anon, 1983; Chuapoehuk & Raksakulthai, 1992), bromelain (Chuapoehuk & Raksakulthai, 1992; Handayani, Ratnadewi, & Santoso, 2007; Subroto,

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Hutuely, Haerudin, & Purnomo, 1985), pepsin (Kumalaningsih, 1986), trypsin and chymotrypsin (Chaveesuk, 1991), as well as trypsin and pepsin (Kristianawati, Ibrahim, & Rianingsih, 2014). Subroto et al. (1985) utilized pineapple juice as the source of bromelain in the processing of fish sauce using by-catch fish as raw material. Good quality fish sauce can be acquired by the use of pineapple extract as much as 8% (v/w) with 10-h incubation period. Sangjindavong, Mookdasanit, Wilaipun, Chuapoehuk, and Akkanvanitch (2009) used pineapple core and pineapple peel for producing fish sauce from surimi waste. Handayani et al. (2007) suggested using 15% NaCl to produce sardine (Sardinella lemuru) sauce with the addition of crude protease extracted from pineapple. Use of crude papain has successfully improved quality and accelerated fermentation process of fish sauce (Lopetcharat, Choi, Park, & Daeschel, 2001; Setyahadi, 2013). A better quality of fish sauce made from Sardinella sp. as raw material was obtained by a combination of 12.5% salt and 1.5% papain, producing a nitrogen conversion of 13.63%. The more the salt addition, the lower the protein degradation rate will be. Increasing papain amount will induce higher nitrogen conversion rate and water-soluble protein degradation level in the liquid. High salt addition level seems to inhibit enzyme activity. On the contrary, the reduction of salt addition level will encourage the growth of microorganisms and generate undesirable odor in the fish sauce. Increasing added papain amount promotes the formation of nitrogen compounds, but results in a viscous material due to connective tissue degradation (Anon, 1983). Chuapoehuk and Raksakulthai (1992) prepared oyster sauce by hydrolyzing minced oyster meat using papain or bromelain supplemented with 20% sodium chloride. It was found that 0.7% papain or 0.3% bromelain produced the highest soluble nitrogen in the hydrolysates and showed no significant differences in proximate composition, pH, consistency, and sensory evaluation scores. Pepsin can be used for fish sauce processing in one condition that the pH of fish is brought down into optimum pH for pepsin activity, i.e., pH 2. Salt amount of 15% is considered suitable for generating an optimum condition in preventing the growth of putrefactive bacteria (Kumalaningsih, 1986). The use of trypsin and chymotrypsin to accelerate the rate of fish sauce fermentation processed from herring (Clupea harengus) increased significantly the rate of proteolysis and the amounts of total nitrogen, formol nitrogen, and free amino acids in the fish sauce product. Fermentation period was

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also reduced from 6–12 to 2 months. A significant increase in total nitrogen and free amino acid contents in the end products was observed when enzyme concentration was increased from 0.3% to 0.6%. Supplementation with 0.6% of 25:75 trypsin:chymotrypsin showed the most satisfactory results in terms of total nitrogen, formol nitrogen, and free amino acid contents. The lighter color of herring sauce produced with 0.6% enzyme supplement was preferred to the darker color of the first grade commercially produced fish sauce. There was no significant difference in the preference for aroma and flavor among enzyme-supplemented sauces and the firstgrade commercially produced fish sauce (Chaveesuk, 1991). Acceleration of fish sauce fermentation process was carried out by Kristianawati et al. (2014) by employing proteolytic enzyme addition and salt level reduction with marine catfish viscera as raw material. The addition of trypsin and pepsin with the concentration of 0.3% produced fish sauce with significantly higher yield and enzyme activity values, as well as better sensory performance. The most organoleptically acceptable fish sauce was obtained through processing with 0.3% trypsin, 20% salt addition, and 45-day fermentation period. Ooshiro et al. (1981) examining papain, bromelain, and trypsin for the production of fish sauce from sardines revealed that papain was the most effective for proteolysis. Fermentation with 0.3% papain at 37°C without adjusting initial pH was the best condition for maximum proteolysis. Manufacturing fish sauce from anchovy fish using purified protease from A. oryzae was investigated by Man and Tuyet (2006). The application of that protease with a suitable procedure of salt addition in fish sauce processing accelerated fish proteolysis and increased the free amino nitrogen content. It should be taken into consideration that high salt content (25%) in the fish–salt mixture decreased the enzyme activity.

5. CONCLUDING REMARKS Fermented fish products play an important role in daily life of many countries. These products are consumed in small amount, but can give a taste sensation-promoting appetite for eating. Endogenous fish proteases, particularly the ones coming from digestive tracts, are suspected as the main enzymes contributing in degrading protein during fermentation process. Eviscerating by removing the gut is not recommended if faster fermentation to be achieved. Microorganisms through excreting proteases seem to take part in the processing of fermented fish mainly either at the beginning of

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the process or prior to the fish receiving salting treatment. In order to obtain optimum involvement of microorganisms in the production of fermented fish, manipulation of the environment to facilitate the most suitable conditions for the growth of microorganisms should be performed. The fermentation process with correct proteolysis is needed to guarantee a good quality product, but this usually requires a long fermentation period, even up to a year. Economically, the shorter fermentation process is more profitable. Therefore, the accelerated processing by enzymes addition has a good prospect as long as the quality of the product is comparable with that obtained without employing enzymes. Intensive studies have been carried out to accelerate fermentation process of fish sauce using the addition of various enzymes manufactured from plants, animals, and microorganisms. Processing acceleration for other fermented products than fish sauce by involving enzyme supplementation is encouraged to be done. Manipulation of fermentation environment to achieve optimum enzyme activities and microorganism growths for speeding up the processing rate of fermented fish products is recommended.

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