Marine Enzymes Biotechnology: Production and Industrial Applications, Part I - Production of Enzymes [1st Edition] 9780128039106, 9780128038475

Table of contents :
Content:
Series PagePage ii
CopyrightPage iv
ContributorsPages ix-x
PrefacePage xiSe-Kwon Kim, Fidel Toldrá
Chapter One - Metagenomics-Guided Mining of Commercially Useful Biocatalysts from Marine MicroorganismsPages 1-26A.R. Uria, D.S. Zilda
Chapter Two - Utilization of Chitinaceous Wastes for the Production of ChitinasePages 27-46S. Das, D. Roy, R. Sen
Chapter Three - Enzymes from Seafood Processing Waste and Their Applications in Seafood ProcessingPages 47-69V. Venugopal
Chapter Four - Marine Fungal and Bacterial Isolates for Lipase Production: A Comparative StudyPages 71-94H.S. Patnala, U. Kabilan, L. Gopalakrishnan, R.M.D. Rao, D.S. Kumar
Chapter Five - Sequential Optimization Methods for Augmentation of Marine Enzymes Production in Solid-State Fermentation: l-Glutaminase Production a Case StudyPages 95-114T. Sathish, K.B. Uppuluri, P. Veera Bramha Chari, D. Kezia
Chapter Six - Solid-State Fermentation vs Submerged Fermentation for the Production of l-AsparaginasePages 115-135K. Doriya, N. Jose, M. Gowda, D.S. Kumar
Chapter Seven - Production of Enzymes from Marine ActinobacteriaPages 137-151X.Q. Zhao, X.N. Xu, L.Y. Chen
Chapter Eight - Recent Advances in Marine Enzymes for Biotechnological ProcessesPages 153-192R.N. Lima, A.L.M. Porto
IndexPages 193-198

Citation preview

ADVISORY BOARDS KEN BUCKLE University of New South Wales, Australia

MARY ELLEN CAMIRE University of Maine, USA

ROGER CLEMENS University of Southern California, USA

HILDEGARDE HEYMANN University of California, Davis, USA

ROBERT HUTKINS University of Nebraska, USA

RONALD JACKSON Brock University, Canada

HUUB LELIEVELD Global Harmonization Initiative, The Netherlands

DARYL B. LUND University of Wisconsin, USA

CONNIE WEAVER Purdue University, USA

RONALD WROLSTAD Oregon State University, USA

SERIES EDITORS GEORGE F. STEWART

(1948–1982)

EMIL M. MRAK

(1948–1987)

C. O. CHICHESTER

(1959–1988)

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

(1989–1993)

STEVE L. TAYLOR

(1995–2011)

JEYAKUMAR HENRY

(2011–2016)

FIDEL TOLDRÁ

(2016– )

Academic Press is an imprint of Elsevier 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States 525 B Street, Suite 1800, San Diego, CA 92101-4495, United States The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom 125 London Wall, London, EC2Y 5AS, United Kingdom First edition 2016 Copyright © 2016 Elsevier Inc. All Rights Reserved No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. ISBN: 978-0-12-803847-5 ISSN: 1043-4526 For information on all Academic Press publications visit our website at https://www.elsevier.com/

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

CONTRIBUTORS L.Y. Chen School of Life Science and Biotechnology, Dalian University of Technology, Dalian, China S. Das Indian Institute of Technology Kharagpur, Kharagpur, India K. Doriya Indian Institute of Technology, Hyderabad, Telangana, India L. Gopalakrishnan PES University, Bangalore, India M. Gowda NITK-Surathkal, Bangalore, India N. Jose Indian Institute of Technology, Hyderabad, Telangana, India U. Kabilan School of Bioengineering, SRM University, Kattankulattur, Tamil Nadu, India D. Kezia Center for Biotechnology, Andhra University, Visakhapatnam, India D.S. Kumar Indian Institute of Technology, Hyderabad, Telangana, India R.N. Lima Laborato´rio de Quı´mica Org^anica e Biocata´lise, Instituto de Quı´mica de Sa˜o Carlos, Universidade de Sa˜o Paulo, Sa˜o Carlos, Sa˜o Paulo, Brazil H.S. Patnala Indian Institute of Technology, Hyderabad, Telangana, India A.L.M. Porto Laborato´rio de Quı´mica Org^anica e Biocata´lise, Instituto de Quı´mica de Sa˜o Carlos, Universidade de Sa˜o Paulo, Sa˜o Carlos, Sa˜o Paulo, Brazil R.M.D. Rao Indian Institute of Technology, Hyderabad, Telangana, India D. Roy Indian Institute of Technology Kharagpur, Kharagpur, India T. Sathish Bioengineering and Environmental Centre, Indian Institute of Chemical Technology, Hyderabad, India R. Sen Indian Institute of Technology Kharagpur, Kharagpur, India

ix

x

Contributors

K.B. Uppuluri Bioprospecting Laboratory, School of Chemical and Biotechnology, SASTRA University, Thanjavur, India A.R. Uria Research and Development Center for Marine and Fisheries Product Processing and Biotechnology, Central Jakarta, Indonesia P. Veera Bramha Chari Krishna University, Machilipatnam, India V. Venugopal Seafood Technology Section, Bhabha Atomic Research Centre, Mumbai, India X.N. Xu School of Life Science and Biotechnology, Dalian University of Technology, Dalian, China X.Q. Zhao State Key Laboratory of Microbial Metabolism and School of Life Science and Biotechnology, Shanghai Jiao Tong University, Shanghai, China D.S. Zilda Research and Development Center for Marine and Fisheries Product Processing and Biotechnology, Central Jakarta, Indonesia

PREFACE In the last decades, progress on 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, easy adaptation to cold conditions, etc. All these properties make marine enzymes very attractive for new catalytic reactions and, of course, new applications in food and nutrition. In view of this increased interest, Advances in Food and Nutrition Research is publishing three consecutive volumes focused on the topic Marine Enzymes Biotechnology: Production and Industrial Application. This volume 78 corresponds to Part I that is mainly dealing with the production of enzymes from marine sources, next volume 79 will correspond to Part II dealing with the marine organisms producing enzymes, and volume 80 will correspond to Part III dealing with the applications of marine enzymes. This volume brings a variety of chapters reporting the production of enzymes from marine sources. So, this includes the metagenomics approach for discovering novel marine enzymes and the production of enzymes from different marine sources like fishery wastes, algae, sponge, fungi, and bacteria. Reported enzymes include proteases, lipases, amylases, cellulases, xylanases, chitinase, laccase, alkaline phosphatase, transglutaminase, hyaluronidase, acetyl glycosaminidase, glutaminase, asparaginase, and carrageenase, among others. The use of solid-state fermentation vs submerged fermentation for the production of marine enzymes is also discussed in several chapters. This volume presents the combined effort of more than 28 professionals with diverse expertise and background. The Guest Editors wish to thank the production staff and all the contributors for sharing their experience and for making this book possible. SE-KWON KIM FIDEL TOLDRA´ Guest Editors

xi

CHAPTER ONE

Metagenomics-Guided Mining of Commercially Useful Biocatalysts from Marine Microorganisms A.R. Uria1, D.S. Zilda Research and Development Center for Marine and Fisheries Product Processing and Biotechnology, Central Jakarta, Indonesia 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Metagenome Isolation 3. Metagenomic Library Construction 4. Functional Screening 5. Homology Screening 6. Conclusions References

1 7 9 12 16 19 21

Abstract Marine microorganisms are a rich reservoir of highly diverse and unique biocatalysts that offer potential applications in food, pharmaceutical, fuel, and cosmetic industries. The fact that only less than 1% of microbes in any marine habitats can be cultured under standard laboratory conditions has hampered access to their extraordinary biocatalytic potential. Metagenomics has recently emerged as a powerful and well-established tool to investigate the vast majority of hidden uncultured microbial diversity for the discovery of novel industrially relevant enzymes from different types of environmental samples, such as seawater, marine sediment, and symbiotic microbial consortia. We discuss here in this review about approaches and methods in metagenomics that have been used and can potentially be used to mine commercially useful biocatalysts from uncultured marine microbes.

1. INTRODUCTION Marine microorganisms play various ecological roles in the oceans, including their involvement in nearly all primary and secondary production as well as in all biogeochemical cycles (Strom, 2008). The Census of Marine Advances in Food and Nutrition Research, Volume 78 ISSN 1043-4526 http://dx.doi.org/10.1016/bs.afnr.2016.05.001

#

2016 Elsevier Inc. All rights reserved.

1

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A.R. Uria and D.S. Zilda

Life program between 2000 and 2010 estimated that the diversity of marine microbes accounts for 1 billion with
38,000 species and 5000–9000 species of microbial bacteria in a liter of seawater and a gram of sand, respectively (Costello et al., 2010; www.coml.org, 2010). The great microbial diversity and vital ecological roles suggest that marine microorganisms are a rich reservoir of highly diverse and unique biocatalysts, either as novel enzymes encoded on discrete single genes or as modular enzymes encoded on gene clusters (Ferrer, Golyshina, et al., 2005; Ferrer, Martinez-Abarca, & Golyshin, 2005). Biocatalysts have attracted much attention for various applications in food, pharmaceutical, fuel, and cosmetic industries, because they are considered as environmental friendly, economical, and clean catalysts that can produce a wide variety of chemical substances (Ferrer, Golyshina, et al., 2005; Ferrer, Martinez-Abarca, et al., 2005). The high efficiency and stereoselectivity of enzyme-mediating reactions make biocatalysts suitable for being applied in various industrial processes, ranging from food-related conversion, laundry detergent, and paper processing to the production of useful fine-chemicals and research reagents (Arnold, 2001; Wahler & Reymond, 2001). For such reasons, it is not surprising that the industrial demand for novel enzymes to improve existing or to establish novel processes has increased dramatically in the past few years (Schmid et al., 2001). Recent particular industrial interests have been focused on novel enzymes capable of mediating new chemical reactions that are difficult or expensive to be achieved by chemical catalysis or traditional organic synthesis (Ferrer, Beloqui, Timmis, & Golyshin, 2009). The general approach of obtaining appropriate microbial biocatalysts with desired specific properties is screening of natural diversity for hitherto unknown enzymes based on traditional cultivation-dependent microbiological methods. This classical approach includes enrichment of microorganisms from environmental samples, isolation of pure cultures, screening of microbial isolates expressing desired enzymatic activity, characterization of the isolated enzymes of interest, and optimizing the enzyme production and recovery (L€ammle et al., 2007). Recent advances in DNA sequencing technologies have contributed to the huge accumulation of new fully sequenced genomes from bacteria and archaea available in the public databases. Examining genome database, popularly called genome mining, provides exciting opportunities of discovering genes encoding novel commercially valuable enzymes with interesting properties (Lee et al., 2010; Lόpes-Lόpes, Cerda´n, Siso, 2014). However, this cultivation-based

Metagenomics-Guided Mining

3

discovery of novel enzymes has so far been limited to only a tiny fraction of the existing microbial diversity in any given habitat, because less than 1% of microorganisms in most environments can be cultivated by conventional methods, as laboratory cultivation conditions do not reflect natural living conditions (Amann, Ludwig, & Schleifer, 1995). The limitation to cultivate most microorganisms under standard laboratory conditions can be overcome by metagenomics, a powerful cultivationindependent tool for analysing the genome sequences of an uncultured microbial community in any given environment (Hugenholtz & Tyson, 2008). In recent years, metagenomics has emerged as a well-established approach to exploit the biocatalytic potential of the vast majority (>99%) of hidden uncultured microbial diversity, leading to the discovery of novel proteins (Lorenz & Eck, 2005). This approach, also called metagenome mining, has been developed to examining metagenome (the collective genomes of all microorganisms present in a given habitat) for the discovery of novel biocatalysts without cultivating any particular microbes (Ferrer et al., 2009; Handelsman, Rondon, Brady, & Clardy, 1998; Steele, Jaeger, Daniel, & Streit, 2009). Metagenomics-guided searching for enzymes from uncultured marine microbes involves direct isolation of total DNA from an environmental sample followed with cloning of the DNA obtained into an easily culturable microbial host. The resulting clones collectively termed as a metagenomic library can be screened to obtain genes of interest (Handelsman, 2004; Handelsman et al., 1998). Further gene sequence analysis, overexpression and characterization of the expression product can provide useful information on the enzyme properties. Approaches for metagenomic library screening can be divided into two general categories, namely, functional screening and homology screening. Functional screening relies on the heterologous expression of the cloned genes in a cultured host to generate phenotypes of interest. Homology screening relies on the use of DNA sequence similarity to identify clones harboring genes of interest in a metagenomic library (Iqbal, Feng, & Brady, 2012). Development of metagenomics techniques has allowed the discovery of an unlimited pool of new potential biocatalysts, genes, and biosynthetic pathways from uncultivable marine microorganisms (Arnold, 2001; Daniel, 2001). Some examples of industrially relevant enzymes discovered from marine microorganisms using metagenomics strategies were listed in Table 1. We describe here common steps in metagenomics that can be used for discovery of marine biocatalysts (Fig. 1), which include metagenome isolation, metagenomic library construction, library screening

Table 1 Examples of Biocatalysts Discovered from Uncultured Marine Microorganisms Through Metagenomics Marine Screening Insert Biocatalyst Source Vector Approach Size (kp) Σ Clone + Clone References

Chitinases

Coastal seawater

Phagemid

Homology

1.8–4.2

75,000

14

Cottrell, Moore, and Kirchman (1999)

Coastal and estuarine water

Plasmid

Homology


0.9

850,000 1

Cottrell, Wood, Yu, and Kirchman (2000)

Sargasso Sea

Plasmid

PCR Homology


0.9

–

108

LeCleir, Buchan, and Hollibaugh (2004)

Amylase

Marine sediment

Fosmid

–

30–40

20,000

1

Liu et al. (2012)

Laccase

Surface seawater

BAC

Homology

50–150

20,000

1

Fang et al. (2011)

Esterases

Deep-sea hypersaline basin

Λ-Phagemid Functional

ND

ND

5

Ferrer, Golyshina, et al. (2005) and Ferrer, Martinez-Abarca, et al. (2005)

Deep-sea sediment Fosmid

Functional


40

ND

1

Park et al. (2007)

Surface seawater

BAC

Functional


70

20,000

4

Chu, He, Guo, and Sun (2008)

Arctic sediment

Fosmid

Functional

–

–

2

Jeon, Kim, Kang, Lee, and Kim (2009)

Sea sediment

Fosmid

Functional


36

–

1

Fu et al. (2011)

Deep-sea sediment Fosmid

Functional


36

20,000

12

Jiang et al. (2012)

Marine sediment

Functional

1–8.5

118,000 15

Peng et al. (2011)

Plasmid

Tidal flat sediment Fosmid

Functional


35

386,400 4

Lee, Lee, Oh, Song, and Yoon (2006)

Deep-sea sediment Fosmid

Functional

21–40

8823

1

Jeon et al. (2009)

Baltic sea sediment Fosmid

Functional

24–39

>7000

1% of library

Ha˚rdeman and Sj€ oling (2007)

Deep-sea sediment Fosmid

Functional

15–33

81,100

6

Jeon et al. (2011)

Marine sediment

Functional

1–8.5

118,000 1

Peng et al. (2014)

Antarctic seawater –

Genome walking

–

–

1

Acevedo et al. (2008)

Deep-sea sediment Fosmid

Functional

ND

ND

1

Lee et al. (2007)

Amidases

Marine sediments

Plasmid

Functional


5.2

25,000

6

Gabor, de Vries, and Janssen (2004)

Cellulase

Sargasso Sea

Fosmid

Homology (WGS)

–

–

1

Cottrell, Yu, and Kirchman (2005)

β-Glucosidase Hydrothermal spring

Plasmid

Functional

4–10

ND

1

Schr€ oder, Elleuche, Blank, and Antranikian (2014)

β-1,4SeaweedEndoglucanase associated microbiota

Plasmid

Functional

1.5–7

40,000

11

Martin et al. (2014)

Fumarase

Plasmid

Homology

1–15


50,000 1

Lipases

Proteases

Seawater

Plasmid

Note: WGS, whole-genome sequencing; ND, not determined.

Jiang et al. (2010)

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A.R. Uria and D.S. Zilda

Direct extraction

Indirect extraction Seawater

Small and middlesized DNA fragments (1–50 kb)

Marine sponge

Single cells Metagenome cloning

Marine sediment

Large DNA fragments (> 50 kb) Metagenome cloning

MDA Agar plate format

Microplate format

3D format

Metagenome sequencing Indicator media

Hybridization

Pooling-PCR-dilution

Clone sequencing

Single-cell sequencing

Metagenomic datasets

Bioinformatic analysis, gene overexpressionand enzyme characterization

Fig. 1 A general scheme of metagenomics for gaining access to commercially useful enzymes from uncultured marine microorganisms. Metagenome can be extracted directly or indirectly from an environmental sample, either seawater, marine sediment, or microbial consortia of marine invertebrates exemplified by sponge. The isolated metagenome was cloned into a heterologous host, usually E. coli, to create a metagenomic library. The vector used for cloning depends on the DNA fragment length. DNA fragments of less than 8 kb are usually cloned using a plasmid, thereby creating a small insert library. Cosmid or fosmid is used to clone fragments of 25–50 kb, generating a middle-size insert library. DNA fragments of >50 kb can be cloned using bacterial artificial chromosome (BAC) to create a large-insert metagenomic library. MDA, multiple displacement amplification.

Metagenomics-Guided Mining

7

by function, homology-driven screening, and metagenome sequencing. Some approaches described here were applied for terrestrial samples, but they can potentially be used or developed for marine samples.

2. METAGENOME ISOLATION In metagenome preparation, three things should be taken into consideration depending on the requirement for downstream steps: DNA yield, length/size, and purity. Metagenomic DNA from an environmental sample can directly be isolated without prior cultivation of microorganisms or cell isolation. This direct metagenome isolation approach usually relies on detergents and enzymes to break and process samples, followed with phenol/ chloroform extraction and isopropanol/ethanol precipitation. This approach results in relatively high DNA yield, but it usually generates small DNA fragments (1–50 kb) due to the shearing or mechanical forces during the extraction process (Desai & Madamwar, 2007; Xing, Zhang, & Huang, 2012). The DNA obtained using this direct approach can be used for the construction of plasmid-based small insert libraries and cosmid/fosmidbased intermediate insert libraries. An example of direct DNA isolation is based on a lysis buffer containing lauryl sarcosyl and urea at high concentration to break microbial cell walls, as reported by Piel et al. (2004). This method is applicable not only for a marine sponge known to harbor the associated complex symbiotic microbes, but also for fish gut sample and marine sediment sample. This method may involve grinding a sample using liquid nitrogen followed with phenol and chloroform extractions. If a sample contains huge amount of polysaccharides as found in many sponges, the cetyltrimethylammonium bromide (CTAB) treatment can be included for polysaccharide removal prior to isopropanol and ethanol precipitation (Gurgui & Piel, 2010; Hrvatin & Piel, 2007). In metagenomics, it is sometimes very important to obtain larger DNA fragments of more than 50 kb, especially for the purpose of generating large insert libraries based on bacterial artificial chromosome (BAC) vector. Avoiding intensive DNA shearing to get larger DNA fragments can be carried out through mechanical separation of prokaryotic or bacterial cells prior to cell lysis and DNA preparation. The drawback of these indirect isolation approach is the relatively low yield, which is 10–100 times lower compared with the direct approach (Parachin, Schelin, Norling, Radstrom, & GorwaGrauslund, 2010). A simple way to separate prokaryotic cells from a marine

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A.R. Uria and D.S. Zilda

sediment sample is based on low-speed centrifugation as reported by Bakken and Lindahl (1995) and Ha˚rdeman and Sj€ oling (2007). Briefly, a sediment sample is suspended in TE-pyrophosphate buffer and gently agitated, followed with blending in a CTAB buffer and low-speed centrifugation to recover prokaryotic cells (Ha˚rdeman & Sj€ oling, 2007). Nycodenz gradient separation method used by Bertrand et al. (2005) to extract bacterial cells from soil samples might be applicable for any marine sediment sample. For a seawater sample, bacterial and archaeal cells can simply be collected by seawater filtration on a 0.22-μm pore size membrane and subsequent membrane washing to release the cells, as reported by Stein, Marsh, Wu, Shizuya, and DeLong (1996) and Chu et al. (2008). To obtain symbiotic microbial symbionts of marine invertebrates, cell separation from the hosts can be conducted, either by simple squeezing as reported for tunicates by Schmidt et al. (2005) or by homogenization/blending and subsequent differential centrifugation as reported for sponges by Bewley, Holland, and Faulkner (1996). The microbial cells obtained from the environmental sample can subsequently be lysed to recover the DNA through several ways. Simple heating reported by Syn and Swarup (2000) was found to be useful in preparing HMW DNA from uncultivated bacterial cells. This protocol involves treatment of the bacterial cells with NE (NaCl and EDTA) buffer to remove extracellular exopolysaccharides followed with heating the cell suspension at 75°C to allow cell lysis. Further phenol/chloroform extraction, isopropanol precipitation, and ethanol washing steps allow the recovery of HMW DNA (Syn & Swarup, 2000). Enzymatic treatment using combination of lysozyme and achromopeptidase allows efficient cell lysis (Bertrand et al., 2005; Courtois et al., 2003). The enzymatically lysed cells are subsequently treated with proteinase K and N-lauroyl sarcosine, followed with applying a cesium chloride density gradient to purify the DNA (Bertrand et al., 2005). Another method to recover HMW from the isolated bacterial cells is the gel-embedding method, as reported by Liles et al. (2008) for soil microorganisms and Quyang et al. (2009) for bacterial cells from diverse sponges. This method starts with embedding the isolated cell pellet in agarose plugs. This is followed with treatment of the plugs with a buffer containing lysozyme to break the embedded cells and then with a buffer containing proteinase K for protein degradation. Further plug electrophoresis allows DNA fractionation and recovery of over 400-kb fragments (Quyang et al., 2009). The HMW DNA fragments of >30 kb length, either

Metagenomics-Guided Mining

9

as the cell-free form or embedded in low-melting point (LMP) agarose plug can be purified by enzymatic gel extraction using GELase commercially provided by Epicentre or by column-based gel extraction. GELase (Epicentre) contains β-agarose-digesting enzyme, enabling breaking down the carbohydrate backbone of LMP gel into small, soluble oligosaccharides (http:// www.epibio.com). The DNA fragments are then recovered by isopropanol or ethanol precipitation (Gurgui & Piel, 2010). Ion exchange chromatography based on resin, commercially provided by QIAGEN (Hilden, Germany, www.qiagen.com), is designed for the recovery of chromosomal DNA of 20–150 kb in size and has been proven to be useful in obtaining pure HMW metagenomic DNA from the uncultured bacterial cell fraction dominated by Entotheonella, filamentous symbionts of marine sponges (Wilson et al., 2014). This method involves careful cell lysis with lysozyme and binding of all the negatively charged molecules (DNA, RNA, and protein) to the resin containing very little salt. When the salt concentration is gradually increased using a medium-salt buffer, traces of RNA and protein become unbound and nonspecific interaction disrupts, thereby leaving DNA on the chromatography column. By passing a highersalt buffer, the DNA is efficiently eluted from the column, which can be concentrated using isopropanol/ethanol precipitation (Qiagen; Brown, 2010).

3. METAGENOMIC LIBRARY CONSTRUCTION For metagenomic library construction, metagenome obtained from an environmental sample needs to be cloned into a cultured bacterial host, usually Escherichia coli, using an appropriate vector. The type of a vector chosen depends on the length of foreign DNA fragments to be cloned. Cloning of DNA fragments of up to 8 kb is usually based high-copy plasmid and phagemid (Brown, 2010). A wide variety of plasmid vectors are available commercially from suppliers. Small-insert libraries based on plasmid are often sufficient to screen a discrete enzyme-encoding genes (average size of 1 kb). Small insert libraries offers an advantage that most genes are present in the appropriate orientation under the influence of the very strong vector expression signals, suggesting a good chance of being expressed and detected by activity screens (Ferrer et al., 2009). Knietsch, Waschkowitz, Bowien, Henne, and Daniel (2003) reported construction of libraries with small inserts to find a clone expressing alcohol dehydrogenase activity.

10

A.R. Uria and D.S. Zilda

Total DNA retrieved from an environmental sample was partially digested using the restriction enzyme, Bsp1431. The resulting small DNA fragments are ligated with pBluescript SK+, tranformed into E. coli DH5α for clone amplification, and transferred to E. coli ECL707 for insert expression. The expression product was then prepared from the host cells at the stationary phase and tested for alcohol dehydrogenase activity (Knietsch et al., 2003). In a similar way, Tirawongsaroj et al. (2008) constructed a small-insert metagenomic library from Thailand hot spring sediments to screen clones expressing lipolytic activity. The metagenomic DNA partially digested with Sau3AI was subjected to size fractionation. Fractions containing DNA fragments of 1–10 kb were selected and cloned into E. coli using a BamHI-linearized zero-background cloning vector (Tirawongsaroj et al., 2008). Creating large-insert libraries increases the chance of finding genes of interest, because less clone numbers need to be screened. The high-capacity vectors such as cosmid can manage DNA fragments of 30–50 kb. An example of a large metagenomic library based on cosmid was reported by Courtois et al. (2003). To increase the ligation process, the DNA fragments of approximately 50 kb was added with polynucleotide (poly-dT) tails prior to ligation with the cosmid previously linearized enzymatically and tailed with poly-dA. The ligation product was packaged in λ phage particles, which were subsequently used to infect E. coli. The resulting clones were arranged and grown in 96-well microtiter plates, allowing long-time storage of the library in glycerol at 80°C (Courtois et al., 2003). Fragments up to 400 kb can be handled using BAC (Brown, 2010; Kim, Shizuya, de Jong, Birren, & Simon, 1992; Shizuya et al., 1992). Interestingly, using BAC, Rondon et al. (2000) constructed two large metagenomic libraries of >1 Gb-inserts and identified genes for lipase, amylase, and nuclease in the libraries. The main problem in cloning of large fragments is the instability and high potential undesired modifications such as rearrangements and deletions as has been shown by Kim et al. (1992) for complex mammalian genomic DNA inserts. To overcome this problem, it requires a high-capacity vector present in low copy number, which can subsequently be induced to high copy number. Obtaining high yields of the cloned DNA is important for downstream applications such as subcloning, enzymatic restriction, sequencing, and recombination. Fosmid is designed to meet this requirement related to the instability and modification of larger inserts. This is best

Metagenomics-Guided Mining

11

exemplified by the fosmid-based vector pCC1FOS (Epicentre) coupled with E. coli host harboring trfA gene that has been proven to be useful in maintaining the stability of large inserts and reducing potential rearrangements. The F factor system present in pCC1FOS controls the low copy number (1–2 copies per cell). If L-arabinose is added into the growth media, this sugar induces the araC-PBAD promoter/regulator system controling trfA transcription, leading the increased copy number to about 10–50 copies per cell in the presence of an oriV replication origin (Westenberg, Bamps, Soedling, Hope, & Dolphin, 2010; Epicentre; Wild, Hradecna, & Szybalski, 2002). Fosmid pCC1FOS is provided by Epicentre in the dephosphorylated linear form to avoid self-ligation. Prior to ligation with this blunt-ended fosmid, DNA fragments should be repaired at the both termini, because the DNA fragment ends can be fray without phosphate residues during extraction and purification processes. This end repairing step relies on T4 DNA polymerase to blunt the frayed ends by filling the excessed 30 -termini and removing the protruding 30 -termini and terminal phosphotransferase to convert the hydroxyl group at the 30 -termini into a phosphate group (Brown, 2010). In a metagenomic library construction based on this fosmid vector, the repaired blunt DNA fragments of approximately 40 kb are ligated with fosmid and packaged in bacteriophage viral particles that are subsequently used to infect E. coli, thereby generating a complex fosmid library as has been reported for sponge samples (Ueoka et al., 2015; Uria, 2012) (Fig. 2). Metagenomic libraries can be generated in different formats, usually agar plates and microplates, depending on the screening approach to be used. Because library in agar plate format cannot be kept alive for a long time, it can be combined into pools, mixed with glycerol and stored at 80°C for a long time. A library in the microplate format can be added with glycerol and stored in the freezer for being used in later screening. However, constructing a very large metagenomic library (up to 1 millions) either in agar plates or microplates is inefficient in term of labor, space, and consumables because it requires a lot of plates or microtiters, a lot of colony picking and larger space for storage. Hrvatin and Piel (2007) have recently developed an efficient 3D library format containing a very large number of clones (up to 1 million clones) with the clone density of 1000 cfu/mL. This library format allowed its long-time storage in smaller space at the freezer and further faster whole-cell screening.

12

A.R. Uria and D.S. Zilda

Sponge sample

Cell separation Bacterial fraction

DNA isolation

DNA isolation

cos camR

HMW metagenomic DNA Fragment selection Selected metagenomic fragment (~ 40 kb) camR cos Packaging

Ligation Fosmid molecules Metagenomic fragment Catenane Catenane Packaging

Clones harboring circular recombinant molecules Transfection into E. coli Chloramphenicol medium

Clones harboring circular recombinant molecules Transfection into E. coli Chloramphenicol medium

Fig. 2 Construction of a metagenomic library using pCC1FOS fosmid vector. Metagenomic DNA obtained from either a sponge sample or bacterial cells associated with sponge is separated on low-melting gel (LMP) via electrophoresis. Subsequently, DNA fragments of approximately 40 kb are ligated with the fosmid pCC1FOS. The ligation product is packaged in bacteriophage viral particles and used to infect E. coli, thereby resulting in recombinant clones, collectively called a metagenomic fosmid library. Redrawn from Uria, A. R. (2012). Investigating natural product biosynthesis in uncultivated symbiotic bacteria of the marine sponge, Theonella swinhoei. PhD dissertation, University of Bonn, Germany. 218 p.

4. FUNCTIONAL SCREENING Functional screening is generally based on detection of the expression products encoded on the individual genes of interest, and therefore it relies

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on the ability of transformants to express genes of interest (Wahler & Reymond, 2001). The success in this functional screening depends on the compatibility of the cloned genes with the expression machinery of the heterologous host, usually E. coli. The most simple functional screening is the use of indicator media containing a substrate for an enzyme of interest. Positive clones expressing the desired enzyme activities can be identified based on the halo formation or color change on agar plate assays. For example, the presence of one of hydrolase members is visualized with a clearing zone around the positive transformants (Rondon et al., 2000). Using indicator media or agar plate assays offer a major advantage that it is highly scalable, which can be integrated with robotics enabling screening of many thousands clones per day (Suenaga, Ohnuki, & Miyazaki, 2007). Functional screening based on indicator media has been developed to gain access to novel esterases from seawater and marine sediment, as recently reported by Chu et al. (2008), Fu et al. (2011), and Jiang et al. (2012). Chu et al. (2008) prepared HMW DNA from bacterial cells collected by filtration of surface seawater. The DNA fragments of >50 kb was cloned into E. coli EPI300 using a BAC. Functional screening of the resulting BAC library consisting of approximately 20,000 clones led to the isolation of four BAC clones with lipolytic activity. Subcloning of two BAC inserts enabled identification of two ORFs, designated as estA and estB, responsible for lipolytic activity. Overexpression and biochemical characterization of them revealed the remarkable activity of EstB toward p-nitrophenyl esters, suggesting its potential industrial applications (Chu et al., 2008). Jiang et al. (2012) have discovered nine genes coding for novel lipolytic enzymes that shared 56–84% identity at the amino acid sequence level to other lipolytic enzymes in the public database. Briefly, they initially prepared DNA from the deepsediments obtained at the depth of 5886 m with the temperature of 1.5°C. A metagenomic library with the average clone insert size of 36 kb was constructed in E. coli using a fosmid, and functionally screened on Luria–Bertani (LB) agar medium containing 1% tributyrin. They found 12 clones exhibiting lipolytic activity based on the clearing zone formation around colonies. Expression and characterization of one of the discovered genes, designated as Est6, revealed that its expression product was an esterase with the preference for mid-length acylglycerols, suggesting its potential industrial application (Jiang et al., 2012). In a separate study by Fu et al. (2011), a metagenomic library constructed from a South China Sea Sediment core was screened for clones exhibiting

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lipolytic activity using the tributyrin agar diffusion assay. This led to the isolation of a positive clone harboring a 36-kb insert fragment. Subcloning of the insert and subsequent sequencing of the positive subclones enabled identification of an esterase gene, designated as EstF. Further phylogenetic analysis, overexpression, and characterization revealed that EstF was active at very low temperatures with the preference for the short acyl chain substrate, suggesting it was a cold-active “true” esterase (Fu et al., 2011). A lipase gene, designated as Est_p6, was isolated by Peng et al. (2014) through functional screening of a marine sediment metagenomic library. Subsequent gene overexpression and protein characterization revealed that this lipase showed a high hydrolytic specificity toward myristate and palmitate as well as a distinctive and desirable flavor and odor in milkfat flavor production, suggesting its potential application for commercial lipolyzed milkfat flavor production and manufacturing processes (Peng et al., 2014). Lipases have been applied in various industries for food, cosmetics, pharmaceutical, chemical, and detergent productions (Kim et al., 2007). Lee et al. (2006) constructed a metagenomic fosmid library from tidal flat sediment and screened for lipolytic activity using LB media containing emulsified tricaprylin. This led to the identification of four clone exhibiting lipolytic activity as shown by a halo surrounding the colonies. Phylogenetic analysis indicated that enzymes expressed by the clones belong to a new family of bacterial lipases (Lee et al., 2006). Functional screening of a metagenomic library for clones capable of secreting cellulase can be based on a colorimetric assay. This assay involves the use of carboxymethylcellulose in the growing media and staining with the dye congo red (Kasana, Salwan, Dhar, Dutt, & Gulati, 2008). The dye interacts with β-D-glucans resulting in the dye–glucan complex indicated with the formation of a yellow halo surrounding of cellulase is the clones expressing cellulase activity (Teather & Wood, 1982). Cellulase has attracted increasing attention in recent years due its great future potential in biofuel production through biodegradation of plant biomass containing lignocellulose (Maki, Leung, & Qin, 2009). Functional screening based on chromogenic or fluorogenic substrates has been used to detect clones expressing certain enzymes that convert the substrates selectively into products which can visually be detected in vivo (Wahler & Reymond, 2001). This screening approach has successfully been developed to identify oxidoreductases, which is applicable for marine metagenomic libraries. For example, clones expressing dioxygenase activity can be screened using horseradish peroxidase (HRP). In this screening, a fluorogenic aromatic substrate is hydrolyzed

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by oxygenases to form a phenol or catechol that is further polymerized by HRP, thereby producing a fluorescent dimer/polymer readily detected spectroscopically using a digital imaging system or plate readers (Joo, Arisawa, Lin, & Arnold, 1999). Another example is the identification of multicopper oxidase in a metagenomic library based on the formation of a catechol (reddish brown) resulted from the guaiacol hydrolysis catalyzed by this oxidase (Ype, Li, Liang, & Liu, 2010). The major disadvantage of indicator media approach is that it relies on transport of either substrate or enzyme of interest to detect activity, and many potential enzymes might not be simply detected based on colony appearance or color changes in the reaction product (Suenaga et al., 2007). To overcome this limitation, some methods have recently been developed to detect the enzyme-catalyzed reaction products. For example, Rabausch et al. (2013) developed functional screening based on a semiautomated thin-layer chromatography (TLC) to screen clone pools in a metagenomic library, allowing the rapid identification of clones expressing flavonoid-modifying enzymes. Clone pools were transferred to a biotransformation medium containing flavonoid, and the reaction products were subjected to TLC analysis. Positive pools were downsized until the responsible single clones were detected (Rabausch et al., 2013). Flavonoid-modifying enzymes can mediate the regio- and stereochemical modification of flavonoids, and therefore they have gained increasing demand for producing specific flavonoids required in the cosmetic, pharma-, and nutraceutical industries (Das & Rosazza, 2006). Another method to overcome such problem is the use of a reporter gene assay called SIGEX (substrate-induced gene expression screening) designed by Uchiyama, Abe, Ikemura, and Watanabe (2005) to screen clones expressing enzymes involved in catabolism. In this method, green fluorescent protein (GFP) was used as reporter to detect metagenomic clones expressing desired catabolic enzymes induced by suitable substrates. SIGEX has allowed discovery of genes involved in benzoate and catechol degradation in a groundwater metagenomic library (Uchiyama et al., 2005). Subsequently, PIGEX (product-induced gene expression) was developed by Uchiyama and Miyasaki (2010) to screen amidase activity in a wastewater sludge metagenomic library. In this method, GFP gene was placed under the control of benzoate-sensitive transcription factor. Amidases expressed by positive clones convert benzamine into benzoate. The resulting benzoate induces the activation of the transcription factor leading to the expression of GFP (Uchiyama & Miyasaki, 2010). In more recently, Hollfelder and his

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colleagues reported an impressive ultrahigh-throughput functional screening for new hydrolases from various metagenomic sources (Colin et al., 2015). This approach was based on the use of microfluidic droplets, in which E. coli cells transformed with environmental DNA using a plasmid were individually encapsulated into single water-in-oil droplets containing substrate and lysis agents. Subsequently, emulsion droplets were incubate offchip and assayed for catalytic activities of cytoplasmically expressed hydrolases after cell lysis (Colin et al., 2015).

5. HOMOLOGY SCREENING Homology-driven screening can be based on PCR using degenerate primers designed specifically to target genes coding for new members of a known enzyme family in an environmental sample. Cloning and sequencing of the PCR product resulted from the degenerate primers can provide insight into the sequence diversity of partial genes with the same function. The sequence information of an amplified partial gene can then be used as the basis for designing new specific primers to recover an entire gene from a metagenomic library. Subsequent heterologous expression of the entire gene may lead to the discovery and overproduction of the encoded biocatalyst. This PCR-based screening is best exemplified by identification of a laccase from the surface water of the South China Sea, as has recently been reported by Fang et al. (2011). The DNA fragments of 50–150 kb prepared from the surface water were cloned into E. coli using a BAC. The resulting clones placed in 385-well plates were screened using degenerate primers designed based on the conserved region of laccases in copper-binding sites. Subsequently, the full-length laccase gene obtained was expressed using the pET expression system. Interestingly, the resulting recombinant laccase (439 amino acids) harboring three conserved copper-binding domains shares less than 40% of sequence identities with known bacterial multicopper oxidases. The unusual properties possessed by this new laccase, such as alkalescence-dependent activity, high chloride tolerance, and dye decolorization ability, makes it an alternative for specific industrial applications (Fang et al., 2011). PCR-based identification of chitinase genes in a seawater sample reported by Cottrell et al. (2000) involved the use of the degenerate primers designed based on deduced amino acid sequences of chitinases in four γ-proteobacteria (Alteromonas sp., Aeromonas caviae, Serratia marcescens, Enterobacter agglomerans). First, bacterial cells were collected from seawater

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samples using 0.22-μm filtration cartridge. Subsequently, PCR amplification was performed using the degenerate primers with total DNA from such bacteria cells as the template. The resulting PCR products were cloned into E. coli, and the resulting clones were selectively sequenced (Cottrell et al., 2000). This PCR-based screening has also been used to identify alkaline proteases as reported by Niehaus et al. (2011). Homology screening based on in situ hybridization can also be used to identify biocatalyst-encoding genes of interest, as shown by Ginolhac et al. (2004) for identifying polyketide synthase in a metagenomic library. This begins with designing degenerate primers to isolate partial genes of interest. The obtained gene fragments are labeled with [α-33P]dCTP and used to screen a metagenomic library. In library screening, colonies on plates are initially transferred on to nylon membranes and lysed to expose the DNA. The DNA on membranes is immobilized and treated with probe to allow visualization of hybridization signals as an indicator of positive clones on a master plate (Ginolhac et al., 2004). Homology-based screening of a metagenomic library constructed from Pacific deep-sea sediment led to the identification of two novel alkane hydroxylases (Xu, Xiao, & Wang, 2008). A very large metagenomic library can effectively be screened using pool dilution and whole-cell PCR method if the library is generated in 3D format (Hrvatin & Piel, 2007) as mentioned in Section 3. In this screening method (Fig. 3), a library is initially grouped into pools with the clone density of 1000 cfus/mL. Then, a small aliquot of several pools is combined as a superpools. Superpools are screened by whole-cel PCR, and the pools from a PCR-positive superpool are subsequently screened. Positive clone pools are diluted at lower cell density, followed by PCR-rescreening. This cell dilution coupled with PCR screening is carried out until the clone density of approximately 20–40 cfus/mL. Cell aliquot with such clone density is then plated at around 200 cfus/plate. Some colonies can be checked by colony PCR to isolate a single positive clone (Hrvatin & Piel, 2007; Uria, 2012). Recent advances in DNA sequencing technologies marked with the development of next-generation sequencing (NGS) has contributed significantly to a great accumulation of metagenomic sequences from various environmental samples. Subsequent bioinformatic analysis of metagenomic datasets allows identifying genes encoding commercially useful enzymes (CUEs). Obtaining the complete sequence of a genome from complex metagenomic DNA is very challenging to accomplish since the metagenomic data come from heterogenous microbial communities, often

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Library screening Production of recombinant biocatalyst

A positive pool (~4000 cfus/mL) + PCR

4 x dilution

Gene cloning and overexpression

Subpools (~1000 cfus/mL)

x 10 +

5 x dilution

PCR-amplification of a target gene

PCR Sub-subpools (~200 cfus/mL)

x 10 +

10 x dilution

Subcloning and Sequencing

PCR Sub-sub-subpools (~20 cfus/mL)

x 10 +

Plating

PCR

Construct preparation

Fig. 3 An example of mathematical calculations for the high-throughput screening of a fosmid-based complex 3D metagenomic library. In an attempt to screen a complex library at high clone density (for instance
4000 cfus/mL), a small aliquot of a positive pool is initially screened by PCR, followed by the combination of pool dilution and whole-cell PCR analysis until the clone density of
20 cfus/mL. Cell suspension is then plated at the clone density of
200 cfus/plate, followed with colony PCR to isolate a single positive clone.

containing more than 10,000 species (Wooley, Godzik, & Friedberg, 2010). In NGS technology, metagenome is usually fragmented into small fragments, generating millions of short reads, commonly in the range of 75–1000 bp depending on the sequencing platform used. The short sequence reads must be assembled to generate contigs, usually up to 5000 bp. Due to the small reads and low depth of coverage, generating accurate and high-quality genome assembly becomes difficult. Although reconstruction of an entire genome present in a complex metagenomic sequences is generally not possible (Riesenfeld, Schloss, & Handelsman, 2004; Wooley et al., 2010), an incomplete metagenomic dataset consisting of contigs assembled from short sequence reads can still offer great opportunities of mining novel functional genes, new activities and products (Ferrer et al., 2009). For example, using the Sargasso Sea whole-genome sequence (WGS) data set prepared by Venter et al. (2004), Cottrell et al. (2005) identified genes coding for hydrolases potentially used by marine Cytophage-like

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bacteria to degrade biopolymers in HMW dissolved organic matter (DOM). PCR primers were then designed to amplify an entire hydrolase gene, designated celM, from a fosmid library constructed with prokaryotic DNA of western Arctic Ocean. Subsequent subcloning of the gene into E. coli and expression product analysis revealed that it has peptidase activity (Cottrell et al., 2005). The existing metagenomic datasets available in public databases can be examined and used as the basis for the discovery of novel industrially relevant biocatalysts. Although the DNA samples are not available, discrete genes that are relatively small in size can be chemically synthesized and heterogously expressed to allow functional analysis, as shown by Bayer et al. (2009) for methyl halide transferase. Samar, Kumar, Prakash, and Taylor (2010) developed MetaBioMe, a database to explore CUEs in metagenomic datasets. Using homology-based approach, they identified several novel homologues for known CUEs, providing exciting opportunities for further experimental verification. Recently single-cell genomics offers great potential to be developed for enzyme discovery. This technique involves isolation of single cells from complex microbial consortia, either by a fluorescence-activated cell sorter as has been shown for bacterial symbionts of a sponge (Unson, Holland, & Faulkner, 1994), microfluidic chip (Marcy et al., 2007), or micromanipulation as reported by Grindberg et al. (2011) for single cells of Lyngbya bouillonii. Subsequent cell lysis and multiple displacement amplification (MDA) to amplify whole genome (Dean et al., 2002; Grindberg et al., 2011) enable obtaining sufficient genomic DNA of an uncultured bacterial species for sequencing. The MDA method relies on the φ29 DNA polymerase coupled with random primers to amplify the entire genome. This polymerase has special characteristics, such as its extremely high processivity and its strong strand displacing activity, which allow copy over the sample template multiple times (Lasken, 2009). Bioinformatic analysis of the MDA-amplified datasets opens exciting chances to identify potential functional genes in an uncultured bacterial species.

6. CONCLUSIONS The enormous abundance and diversity of marine microorganisms offer exciting opportunities and challenges in discovery of highly diverse and unique novel biocatalysts that are commercially useful or industrially relevant. The fact that less than 1% of microorganisms in any given environment cannot be cultivated under traditional laboratory conditions has

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hampered exploring and exploiting the vast majority of marine microorganisms. Metagenomics has recently emerged as powerful and well-established tool to access the biocatalytic potential of uncultured microbial communities living in marine habitats, leading to the discovery of novel enzymes with industrial potential applications. Metagenomics-guided mining of marine microorganisms for CUEs involves direct or indirect extraction of total DNA from an environmental sample (called metagenome) followed with metagenome cloning, thereby resulting in a metagenomic library. Libraries can be generated in different formats, either agar plates, microplates, or 3D semiliquid, depending on the library size and the screening approach to be used. Subsequent library screening, gene analysis, and overexpression may lead to the discovery of novel recombinant enzymes that can be produced at the large scale in sustainable way. Further enzyme characterization provides useful information on the enzyme properties that are important for industrial applications. Approaches for metagenomic library screening can be divided into two general categories, namely, functional screening and homology screening. Functional screening relies on the heterologous expression of the cloned genes in a cultured host to generate phenotypes of interest. This screening approach includes the use of indicator media containing a substrate for an enzyme of interest to identify clones expressing the desired enzyme activities based on the halo formation or color change on agar plate assays. Another functional screening is based on the use of chromogenic/fluorogenic substrates to identify clones expressing certain enzymes capable of converting the substrates into in vivo-detectable products. Many potential enzymes might not be simply detected based on indicator media or chromogenic/ fluorogenic substrates. To overcome such limitation, other methods have developed to detect the reaction product catalyzed by the enzymes of interest, exemplified by automated TLC and using the reporter gene encoding GFP placed under the transcription factor. Homology-driven screening is usually based on PCR or hybridization. Especially, the pool dilution and whole-cell PCR method developed by Hrvatin and Piel (2007) allows faster screening of a highly complex metagenomic library. Recent advances in DNA sequences technologies have enabled fast sequencing of the metagenome come from a uncultured bacterial community, leading to the accumulation of metagenomics datasets. Single-cell genomics has recently emerged as a powerful tool to examine a genome from an uncultured single bacterial species. Bioinformatic analyses of metagenomic datasets open exciting chances to identify potential novel genes encoding CUEs.

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CHAPTER TWO

Utilization of Chitinaceous Wastes for the Production of Chitinase S. Das, D. Roy, R. Sen1 Indian Institute of Technology Kharagpur, Kharagpur, India 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Chitinaceous Wastes and Chitin 2.1 Chitin Structure and Complexes 2.2 Chitin Sources 3. Chitinase 4. Chitinase Production 4.1 Submerged Fermentation 4.2 Solid-State Fermentation 4.3 Fed-Batch Fermentation 4.4 Continuous Process 4.5 Biphasic Cell System 4.6 Recombinant Technology 5. Conclusions Acknowledgments References

28 28 29 31 32 34 35 37 38 40 40 41 41 42 42

Abstract Marine environment is the most abundant source of chitin. Several marine organisms possess chitin in their structural components. Hence, a huge amount of chitin wastes is deposited in marine environment when such organisms shed their outer skeleton and also after their demise. Waste chitins are potential nutrient source of certain microbes. These microbes produce chitinases that hydrolyze waste chitins. These organisms thus play an important role to remove the chitin wastes from marine environment. In connection with this, chitinases are found to be most important biocatalyst for the utilization of chitin wastes. Therefore, use of chitin for chitinase production is one of the useful tools for different types of bioprocesses.

Advances in Food and Nutrition Research, Volume 78 ISSN 1043-4526 http://dx.doi.org/10.1016/bs.afnr.2016.04.001

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2016 Elsevier Inc. All rights reserved.

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1. INTRODUCTION Chitin is a second most abundant natural biopolymer after cellulose. It is a homopolymer of N-acetyl-D-glucosamine (GlcNAc) linked by β-(1,4) bond. It is distributed in nature as structural body parts of fungi, marine diatoms, mollusks, arthropods, crustaceans, nematodes, etc. (Chen & Lee, 1995). Large amount of waste chitin is deposited in food processing and enzyme production industries; and removal of the waste chitin from different industries becomes a serious issue. Conventional method for chitin disposal by burning, land filling, and ocean dumping suffer from several environmental and economic issues (Suresh & Chandrasekaran, 1998). On the other hand, demands on chitin degrading enzyme, chitinase, production is increased from last few decades due to its potential biotechnological applications in pharmaceutical and biofuel industries (Dahiya, Tewari, & Hoondal, 2006). Microbial enzymes are observed to be advantageous over the enzyme from higher organisms due to low cost, variability in catalytic activity, and higher stability (Chandrasekaran, 1997). So, exploitation of waste chitin for microbial chitinase production by different bioprocesses is thought to be an important aspect for waste chitin utilization and bioremediation (Krishnaveni & Ragunathan, 2014).

2. CHITINACEOUS WASTES AND CHITIN It is estimated that out of total wastes generated in food processing industry around 20–50% are comprised of chitin (Wang & Chang, 1997). These are mostly generated during shell fish processing. Therefore, shellfish are treated as good source of chitin. But, there are certain limitations in chitin extraction from shellfish wastes—such as, seasonal harvesting of crustaceans, limited raw material supply, and generation of environmental hazardous material during deproteination of waste chitin. These are the reasons why the researchers are striving to search for new and sustainable sources of chitin. Fungal biomass may emerge as a potentially viable and alternative source for chitin production, since fungal cell wall is composed of mainly chitin biopolymer (Wang & Chang, 1997). Moreover, cost effective fungal cultivation in solid state and submerged cultures would be an added advantage to obtain required amount of chitin. On the other hand,

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Chitinase Production by Using Waste Chitin

chitin can be obtained from pupa silkworm as reported in recent past (Haga & Shirata, 1997). Following sections are dealt with chitin structure, complexes, and its availability from different sources.

2.1 Chitin Structure and Complexes Chitin is mainly composed of N-acetyl-D-glucosamine residues, which are attached by β-(1,4)-glycosidic linkage (Fig. 1). Important functional groups of chitin are listed in Table 1. Chitin is recognized as widely distributed in exoskeleton of invertebrates and cell wall of fungi. Chitin and cellulose are structurally analogous, where in the former the hydroxyl group of glucose moiety is replaced by the acetamide group. However, degree of acetylation

Fig. 1 Linear structure of chitin.

Table 1 List of Functional Groups Inferred from FTIR Inference from Spectral Response Functional Group (Mudasir, Tahir, & Wahyni, 2008)

dOH

dOH stretching (“a,” Fig. 1)

dCH3CONH

dNH stretching (“b,” Fig. 1) dCH stretching (“c,” Fig. 1) ]CO stretching (“d,” Fig. 1) dNH bending (“b,” Fig. 1) dCH bending (“c,” Fig. 1) dCN stretching (“e,” Fig. 1)

]CO of polysaccharides and glucosamine rings

]CO stretching (“f,” Fig. 1)

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in the polysaccharide molecule depends on the source and chitin isolation process. At least, one out of six GlcNAc molecules remains deacetylated in native chitin (Trudel & Asselin, 1990). It is observed that length of the yeast chitin is only with 100 GlcNAc residues, whereas the crab chitin contains 5000–8000 GlcNAc residues. Chitin polymer is labile to alkali. It was observed that the length of the chitin chain was decreased upon hot alkali treatment during the deproteinization step of chitin isolation (Synowiecki & Al-Khateeb, 2003). Each polymeric chitin chain is associated with neighboring chain by hydrogen bond, where amino group (>NH) of one molecule makes bond with carbonyl (>C]O) group of the adjacent one. On the basis of nature of H-bond, chitin molecule can be designated as α, β, and γ chitin. In α-chitin, the chains are arranged in parallel fashion ("") and it is mostly found in arthropods and crustaceans; whereas, an antiparallel arrangement of chitin chain is observed in β-chitin ("#) and these are obtained from marine diatoms. In contrast, in γ chitin, the arrangement of the chitin chain is little complicated; where, out of each three chitin chain, two are arranged in parallel fashion and third one is arranged in antiparallel fashion (""#). However, the existence of γ chitin is a matter of controversy (Chanzy, 1997). The α-chitin is observed to be most stable form of chitin; while β-chitin can easily be converted to α-chitin by lithium thiocyanate treatment or formic acid precipitation (Hackman & Goldberg, 1965). Hardness, flexibility and permeability of the shell are determined by the ratio of α-chitin and β-chitin. Chitin cannot be melted in solid state due to the presence of high density of hydrogen bonds. However, it can be dissolved in concentrated acids, like hydrochloric acid, sulphuric acid or phosphoric acid, trichloroacetic acid, and formic acid. Greater solubilization rate is observed for β-chitin in comparison to the α-chitin. Besides this, chitin gets solubilized in hexafluoroisopropanol or hexafluoroacetone and some of the hot and concentrated neutral salts (Synowiecki & Al-Khateeb, 2003). Chitin microfibrils are generally resistant to the deacetylases due to the presence of hydrogen bonds. Deacetylases act on the newly synthesized chitin in chitosome prior to synthesis of chitin microfibrils (Kolodziejska & Sikorski, 1999). Several biomolecules like protein, other polysaccharides, and matrix of β-glucan molecules surround the chitin fibril in fungal cell wall to form alkali-insoluble complexes. In crustacean and insects, chitin forms a

Chitinase Production by Using Waste Chitin

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complex with proteins which are tanned by phenolic derivatives. Thus, a glycoprotein framework is constituted in these organisms. Finally, the presence of mineral salts, carotenoids, lipoproteins, and waxes surrounding the glycoprotein frame work, influence the elasticity, permeability, tensile strength, and hardness of the structure. Moreover, calcium carbonate and calcium phosphate, to some extent play a crucial role for the hardness of insect cuticle. Although, silica and iron oxide were also reportedly enhance the hardness of Rhizopods shell (Roberts, 1992).

2.2 Chitin Sources Chitin is widely available from two natural resources such as wastes from marine aqua animal shells and microbes which are described below. Recently, chitin is produced in large scale from the waste of crustacean harvesting industry. The data available on the amount of chitin resources indicate that approximately 0.7, 1.4, and 29.9 million tons of chitin are annually recovered from squid, oyster, and shellfish, respectively (Synowiecki & Al-Khateeb, 2000). Certain crustacean orders (Decapoda, Amphipoda, Copepoda, Anostraca, and Cladocera) are good sources of chitin as they have significant amount (2–12%) of chitin content in their total body mass (Johnson & Peniston, 1982). Crab and shrimp shells, on the other hand are rich source of chitin. In US around 39,000 tons of shell from crab (Cancer magister) and shrimp (Pandalus borealis) is being discarded annually (Knorr, 1991). Moreover, around 50,000 tons of shell is being recovered from the harvested crab (Chionoecetes opilio sp.) in the Canadian Atlantic regions; where, 80% of total weight is generally used to produce the by-products. Shrimp and krill wastes contain around 10% higher chitin than crab-processing wastes (Naczk, Synowiecki, & Sikorski, 1981). A similar amount of chitin is found in Louisiana crawfish (Procambarus clarkii). In crab, chitin is mainly deposited in leg, shoulder, and tips rather than other body parts (Shahidi & Synowiecki, 1991). Antarctic krill (Euphausia superba) is another important chitin source (Kolodziejska, Malesa-Cie´cwierz, Lerska, & Sikorski, 1999). However, fluoride and other minerals are found as a contaminant in krill shell, which occupies more than 10% of the weight (Kolodziejska et al., 1999). Fungal biomass is also known as an abundant source of chitin. It is observed that chitin content varies among the different strains of fungus (Muzzarelli, Ilari, Tarsi, Dubini, & Xia, 1994). The filamentous fungal strains such as Rhizopus, Penicillium, Aspergillus, Fusarium,

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Choanephora, Zygorhynchus are known to be used for chitin extraction and isolation (Muzzarelli et al., 1994). Along with chitin, chitosan and other polysaccharides are also present in the fungal cell wall (Knorr, Beaumont, & Pandya, 1989). In recent years, several attempts were made to isolate chitin from nonconventional fungal resources, because of consistent availability of fungal biomasses from different industries (Muzzarelli et al., 1994). The use of fungal chitin has sometimes advantageous over the shellfish chitin, for high growth rate of fungal strains, inexpensive cultivation in waste materials; absence of high amount of mineral salts and lower pretreatment cost. Besides, it is possible to control the fermentation, processing, and genetic modification of the fungal strains for improving the chitin yield. The chitin content varies from 2% to 60% on the basis of fungal cell wall dry weight. Overall 26–65% and 22–67% chitins and glucans, respectively, have been estimated from deproteinized cell wall of Basidiomycetes and Ascomycetes. For example, Agaricus bisporus, a very commonly known mushroom, is observed to possess proteins (22%), chitin (72%), cellulose (3%) along with microamount of glucosamine, and mineral salts in their cell wall. In mold mycelia, up to 25% protein, 3% nucleic acid, and 15% lipid are found to be present on the basis of their dry weight (Synowiecki & Al-Khateeb, 1997). High chitin content, up to 35% dry cell wall weight has been reported from Mucor rouxii (Bartnicki-Garcia & Nickerson, 1962; Synowiecki & Al-Khateeb, 1997). In addition to this, cell wall of Aphyllophorales is known to possess very high level of chitin content (almost 95%, w/w) in its cell wall (Gorovoj & Kosyakov, 1997). It has been found that it is necessary to optimize the growth medium, time, and other physical parameters for fungal cultivation to maximize the chitin content in fungal cell wall. However, this waste chitin is processed for chitin isolation and they are used for chitinase production.

3. CHITINASE Chitinases (EC 3.2.1.14) are a group of hydrolytic enzymes that cleave the β-1,4-glycosidic bond between two consecutive N-acetyl-Dglucosamines (GlcNAc) of chitin chain (Xia et al., 2001). Wide ranges of organisms are known to produce chitinases, which play diverse role in different organisms which are listed in Table 2. Out of these different sources, microbial chitinases are known to possess potential industrial applications in agricultural and environmental sectors, such as, production of GlcNAc and

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Chitinase Production by Using Waste Chitin

Table 2 Chitinase-Producing Organism and Their Functions Chitinase Producer Requirement to Produce Chitinase References

Virus

Pathogenesis

Patil, Ghormade, and Deshpande (2000)

Fungus

Cell wall remodeling, morphogenesis, Adams (2004) and nutritional purposes

Bacteria

Chitin digestion for nutrition, energy Tsujibo et al. (1993), Park et al. source, and chitin recycling (1997), Svitil, Chadhain, Moore, and Kirchman (1997), and Wang and Chang (1997)

Insect

Postembryonic development and old Merzendorfer and Zimoch cuticle degradation (2003)

Plants

Defense mechanism against pathogen Graham and Sticklen (1994) and plant development

Human

Defense purposes

Boot et al. (2001, 2005) and van Eijk et al. (2005)

other chito-oligosaccharides for healthcare, isolation of fungal protoplasts for strain improvement (Dahiya, Tewari, Tiwari, & Hoondal, 2005), control of pathogenic fungi and mosquito by dissolving cell wall or shell chitin (De Marco, Lima, Desousa, & Felix, 2000; Mendonsa, Vartak, Rao, & Deshpande, 1996), and bioconversion of different chitinaceous wastes (Dahiya et al., 2006). Classification and nomenclature of chitinolytic enzyme are still not well defined. In a contemporary research, Graham and Sticklen proposed two major categories of chitinase—endochitinase and exochitinase (Graham & Sticklen, 1994). A group of chitinases that cleave randomly at internal sites of the chitin chain and produce low-molecular-weight oligomers of GlcNAc like chitotetraose, chitotriose, etc., are known as endochitinase (EC 3.2.1.14). On the other hand, the chitinases which cleave the chitin chain from terminal ends fall under exochitinases. According to the release of the product, exochitinases are again categorized into two subcategories— chitobiosidase and β-(1,4)-N-acetyl-glucosaminidase. An exochitinase, which releases only diacetylchitobioses from the nonreducing end of the chitin, is designated as chitobiosidases (EC 3.2.1.29). In contrast, β-(1,4)N-acetyl-glucosaminidases (EC 3.2.1.30) release only GlcNAc as sole product from nonreducing end of the chitin polysaccharide (Fig. 2).

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S. Das et al.

Fig. 2 Mode of action of different types of chitinases.

4. CHITINASE PRODUCTION Chitinase is produced by number of biotechnological methods, such as submerged fermentation, solid state fermentation, fed-batch fermentation, continuous fermentation, and biphasic cell system, etc. It is observed

Chitinase Production by Using Waste Chitin

35

that expression of extracellular chitinases is induced by external chitin (Dahiya et al., 2006). The organisms producing extracellular chitinases consume chitin as potential carbon and nitrogen source. In this regard, chitin from crab or shrimp wastes could be used in chitinase production media. Chitinase production is found to be greatly affected upon physical and chemical changes (Dahiya et al., 2006). Presence of magnesium sulfate (MgSO4 7H20) and potassium dihydrogen phosphate (KH2PO4) in media has significant effects on the chitinase production (Khan, Hamid, Ahmad, Abdin, & Javed, 2010). However, certain organic supplements and sugar molecules exhibit contradicting effect on chitinase production in different organisms. Yeast extract shows antagonistic effect on chitinase production in Stenotrophomonas maltophilia (Khan et al., 2010); but the same component shows enhancement of chitinase production in Trichoderma harzianum (Nampoothiri et al., 2004). Same observation is found in case of glucose also. Chitinases production increases when glucose is added as supplement with chitin in production media of Enterobacter sp. (Dahiya et al., 2005). However the same glucose imparts a negative impact on chitinase production of Streptomyces lividans as observed by Miyashita, Fujii, and Sawada (1991). Besides, several agroindustrial wastes, such as rice bran and wheat bran serve as nutrient supplements in the media (Dahiya et al., 2005). Addition of amino acid like tryptophan, tyrosine, glutamine, and arginine in medium are shown to enhance the chitinase production of Bacillus sp. as reported previously (Dahiya et al., 2006). Along with chemical parameters, some physical parameters like pH of the production medium, aeration, inoculum size, temperature, and time of incubation also play a crucial role in the chitinase production (Dahiya et al., 2006). Discussion on different methods of chitinase production is a helpful tool for better understanding of the process.

4.1 Submerged Fermentation Selection of cultivation process is an important step for chitinase production. In most of the cases extracellular chitinases are produced in medium at low concentration. Handling of low amount of enzyme in large volume of media is a problem; which complicates further purification steps (Stoykov, Pavlov, & Krastanov, 2015). On the other hand, submerged fermentation (SMF) technique have certain features, such as well-controlled process parameters, increased mass transfer, and enhancement of oxygen delivery system, etc., which help it gaining added advantages over liquid culture (Stoykov et al., 2015). Besides, coimmobilization method has been

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developed to enhance chitinase production. Earlier, calcium alginate was used to coimmobilize the microbial cells and chitin for the production of chitinase from Micromonospora chalcea (O’Riordan, McHale, Gallagher, & McHal, 1989). In this case, chitinase was separately produced by free cells and coimmobilized cells in a production medium with 2% chitin. Coimmobilization helped increase chitinase production by 0.3 U as shown in earlier literature (O’Riordan et al., 1989). Several microbial strains studied for chitinase production by SMF are listed below (Table 3). Table 3 Microbial Chitinase Production by SMF Medium pH, Incubation Substrate Temperature (°C), Microorganism Habitat Used Chitinase Activity (U) References

Nocardia orientalis

Terrestrial Colloidal chitin

5, 28, 1.278

Usui, Hayashia, Nanjob, Sakaib, and Ishido (1987)

Serratia liquefaciens

Terrestrial Chitin

7.5, 30, 15.1

Joshi, Kozlowski, Richens, and Comberbach (1989)

Myrothecium verbucaria

Terrestrial Sclerotium 5.5, 28, 0.24 rolfsii mycelium

Vyas and Deshpande (1989)

Bacillus licheniformis

Terrestrial Collidal chitin

7, 50, 0.014

Takayanagi, Ajisaka, Takiguchi, and Shimahara (1991)

Talaromyces emersonii

Terrestrial Chitin

5, 45, 0.45

James, Hackett, Tuohy, and Coughlan (1991)

Streptomyces cinereoruber

Terrestrial Aspergillus 6.8, 30,17 niger cell wall

Tagawa and Okazaki (1991)

Vibrio alginolyticus

Terrestrial Squid chitin

7, 37, 5.8

Ohishi et al. (1996)

Monascus purpureus

Terrestrial Shrimp and crab shell powder

7, 25, 191

Wang, Hsiao, and Chang (2002)

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Chitinase Production by Using Waste Chitin

Table 3 Microbial Chitinase Production by SMF—cont'd Medium pH, Incubation Substrate Temperature (°C), Microorganism Habitat Used Chitinase Activity (U) References

Trichoderma harzianum

Terrestrial Chitin

5.5, 30, 14.7

Sandhya et al. (2004)

Aeromonas sp.

Marine

5,40, 1.1

Kuk et al. (2005)

Trichothecium roseum

Terrestrial Crab shell 6, 28, 0.78 chitin

Li, Zhang, Liu, and Lu (2004)

Aspergillus fumigatus

Terrestrial Colloidal chitosan

7, 37, 5.86

Jung, Kuk, Kim, Jung, and Park (2006)

Aspergillus terreus

Terrestrial Fish scale waste

6, 30, 4.31

Ghanem, Al-Garni, and Al-Makishah (2010)

7, 10, 0.095

Velmurugan, Kalpana, Hoon, Jung, and Yang (2011)

Plectosphaerella Marine sp.

Swollen chitin

Colloidal chitin

Trichoderma aureoviridae

Terrestrial Colloidal chitin

4.7, 28, 0.036

Agrawal and Kotasthane (2012)

Aspergillus terreus

Marine

6, 30, 4.7

Krishnaveni and Ragunathan (2014)

Shrimp waste

Along with the indigenous chitinase production by microorganisms, statistical methods are employed to optimize production medium constituents and enhancement of chitinase production. Khan et al. in 2010 validated the Design Expert prediction tool to enhance chitinase production in Stenotrophomonas maltophilia. Several reports on chitinase production enhancement by using statistical methodology are listed in Table 4.

4.2 Solid-State Fermentation Use of solid support or matrix during microbial growth and production of metabolites is known as solid state fermentation (SSF). It is commonly used for microbial enzyme production in different sectors especially for fungal chitinase production. Although, SSF experiences some drawbacks, such as

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Table 4 Microbial Chitinase Production by SMF and Statistical Optimization Increase in Chitinase Microorganism Habitat Statistical Method Used Production References

Alcaligenes xylosoxidans

Marine

Plackett–Burman design 2.42-fold and Box–Behnken response surface methodology

Vaidya, Vyas, and Chhatpar (2003)

29%, 9.3%, and 28%

Nawani and Kapadnis (2005)

Pantoea dispersa Marine

Plackett–Burman design 4.21-fold

Gohel, Chaudhary, Vyas, and Chhatpar (2006)

Streptomyces sp. Marine Da11

Plackett–Burman design 39.2-fold and Box–Behnken response surface methodology

Han, Li, Miao, and Zhang (2008)

Streptomyces sp. Terrestrial Response surface methodology and NK1057, numerical optimization NK528, and NK951

uncontrolled process parameters for maintenance of pH, temperature, substrate sterilization, culture purity, and process length; but, the use of cost effective matrices and easy operation of the process make it advantageous over the SMF (Karthik, Akanksha, & Pandey, 2014). In nature, chitin is present as solid and insoluble substrate. Thus, use of chitin from natural resources becomes an additional advantage for chitinase production by SSF. In several literatures different organisms are shown to produce chitinase by SSF. These are listed in Table 5.

4.3 Fed-Batch Fermentation Fed-batch fermentation is alternatively known as semi-batch culture where nutrient or substrates are supplied to the bioreactor according to the required amount. Thus, fed-substrate concentration can be controlled easily (Yamane & Shimizu, 2005). Earlier, in N-acetyl-D-glucosamine production Serratia marcescens QM B1466 was used to express chitinase in fed-batch fermentation (Kim, Creagh, & Haynes, 1998). The culture was grown in 10 L bioreactor for 7 days by using chitinaceous wastes (crab/shrimp chitin) in the media. The pH was set at 8.5 and temperature at 30°C with 3 h feeding time

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Table 5 Microbial Chitinase Production by SSF

Microorganism Habitat

Substrate Used

Medium pH, Incubation Temperature (°C), Chitinase Activity (U/gds)

Beauveria bassiana

Marine

Trichoderma harzianum

Terrestrial Wheat bran 4.5, 30, 3.14 and colloidal chitin

Prawn waste 9.5, 27, 248

Enterobacter sp. Terrestrial Wheat bran and flake chitin

8, 30, 1475

References

Suresh and Chandrasekaran (1998) Nampoothiri et al. (2004)

Dahiya et al. (2005)

Verticillium lecanii

Terrestrial Shrimp 6, 25, 1674 waste silage and sugar cane baggase

Matsumoto, SaucedoCastan˜eda, Revah, and Shirai (2004)

Fusarium oxysporum

Terrestrial Wheat bran and flake chitin

6, 30, 23.6

Gkargkas et al. (2004)

Penicillium aculeatum

Terrestrial Wheat bran and flake chitin

5, 30, 12.53

Binod, Chandran, Pradeep, George, and Ashok (2007)

8.6, 32, 311.84

Suresh, Anil Kumar, and Sachindra (2011) and Suresh, Sachindra, and Bhaskar (2011)

Penicillium Terrestrial Wheat bran monoverticillium and shrimp shell chitin waste Oerskovia xanthineolytica

Terrestrial Wheat bran 7.5, 45, 148 and colloidal chitin

Waghmare, Kulkarni, and Ghosh (2011)

(Kim et al., 1998). Recently, Rao, Inman, Holmes, and Lalitha (2013) also reported the use of fed-batch fermentation for chitinase production in a mixed culture of Vibrio harveyi and Vibrio alginolyticus. In this study, a 10 L bioreactor was used with daily addition of 2% colloidal chitin (w/v) at 30°C, with 20% dissolved oxygen and 150 rpm agitation. With fed-batch fermentation up to threefold chitinase activity can be achieved. Thus, fed-batch fermentation is known to be effective process for chitinase production.

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4.4 Continuous Process Feeding of inputs (raw material, auxillary materials, energy) in a system and extraction of output from the system simultaneously at constant rate and ratio is known as continuous process. Attempts are made by researchers for continuous chitinase production. Fenice, Giambattista, Raetz, Leuba, and Federici (1998) reported continuous chitinase production by immobilized conidiophore of Penicillium janthinellum P9. In this report, polyurethane sponge and chemically modified macroporous cellulose are used as matrix for conidiophore immobilization; where modified cellulose is observed to be better carrier in terms of negligible amount of cell leakage (Fenice et al., 1998). Here, rapid chitinase production is observed by cell-immobilized repeated batch process compare to free cell-based process. Additionally, continuous increase in biomass in the same bioreactor results in the increase of chitinase production in repeated batches. In general, the continuous process is maintained at 168 h. Continuous processes for chitinase production was reported in 1 L fluidized bed reactor where immobilized P. janthinellum P9 were used (Fenice et al., 1998). Unlike repeated batch culture the continuous process generates more chitinase. It was observed that the amount of chitinase from repeated batch culture was 338 U/L, whereas with continuous process the amount of chitinase was increased to 450 U/L. Another mode of continuous chitinase production reported earlier is a membrane-based fermentation process (Kao, Huang, Chang, & Liu, 2007). A strain of Paenibacillus sp. CHE-N1 is used in this study. A membrane outer recycling loop is equipped with the bioreactor; where the effect of membrane pore size is evaluated for different process-related parameters like chitinase recovery, fouling, cellretention efficiency, etc. Here, 300 kDa pores sized M 9 microfiltration column is used for chitinase production. In previous report, colloidal chitin from crab shell wastes was used to feed every 3–4 days which resulted 78% increase in total chitinase activity (42.8 U) compared to batch mode (Kao et al., 2007).

4.5 Biphasic Cell System A PEG/dextran aqueous two-phase system (ATPS) was developed by Chen and Lee (1995) for the chitinase production enhancement by Serratia marcescens. In this process, the production media contains 2% PEG and 5% dextran ATPs. The swollen chitin is used as inducer for chitinase production. A polymer free reference system is used to compare the results

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Table 6 Chitinase Production by Recombinant Microbial Strains Substrate Medium pH, Incubation Temperature (°C), Used/ Chitinase Activity (U) References Inducer Microorganism

Serratia marcescens Chitin designated as BL40

7.2, 30, 184

Kole and Altosaar (1985)

Serratia marcescens SJ101

7.5, 30, 15 Cane molasses and chitin

Joshi et al. (1989)

Recombinant Escherichia coli gene from Aeromonas hydrophila

IPTG

7, 37, 0.903

Chen, Chang, and Cheng (1997)

Self-fusion of protoplasts of Trichoderma harzianum

Chitin

6.5, 25, 9.6

Prabavathy, Mathivanan, Sagadevan, Murugesan, and Lalithakumari (2006)

of fermentation. Here, 2.5%, 1.9%, and 3.1% increase in chitinase activity can be achieved in dextran solution, PEG solution, and ATPS.

4.6 Recombinant Technology Number of wild-type microbial strains is known to produce chitinase. But, hyper-chitinase producer are highly desirable for its different industrial applications. Several fermentation processes are reported to increase productivity by the use of recombinant strains (Karthik et al., 2014). In this regards, some of the recombinant strains are also reported to produce high level of chitinase. Reports on enhanced chitinase production by recombinant strains are listed Table 6.

5. CONCLUSIONS Marine environment consists of three fourth part of the earth surface and it is the richest source of chitinaceous wastes. These are recycled by the action of different microbial enzymes like chitinase. Thus, exploitation of chitin waste is found to be best option for microbial chitinase production. Unfortunately, very few marine microorganisms are reported to utilize waste chitin for chitinase production, therefore discovery marine

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microorganisms is necessary for chitinase production by the utilization of waste chitin in future. Besides, research on large-scale chitinase production is scanty. Detailed investigation on kinetic studies for fermentation, designing of bioreactor, influence of reactor parameters like agitation, aeration is also required for large-scale chitinase production. These investigations may be brought under consideration for future scope for chitinase production research.

ACKNOWLEDGMENTS SD is thankful to National Jute Board (Govt. of India) and HRDG (Govt. of India) for financial support for this study.

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CHAPTER THREE

Enzymes from Seafood Processing Waste and Their Applications in Seafood Processing V. Venugopal1 Seafood Technology Section, Bhabha Atomic Research Centre, Mumbai, India 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 2. Enzymes from Fish Processing Waste 2.1 Fish Proteases 2.2 Lipases 2.3 Transglutaminases 2.4 Carbohydrases 2.5 Other Enzymes 2.6 Recovery of Enzymes from Seafood Waste for Seafood Processing 3. Applications of Fishery Enzymes in Seafood Processing 3.1 Proteases 3.2 Transglutaminase 3.3 Lipases 3.4 Carbohydrases 3.5 Quality Control 3.6 Uses of Cold-Adapted Enzymes 4. Commercial Aspects References

48 48 49 52 52 52 53 53 53 54 57 60 60 61 63 63 64

Abstract Commercial fishery processing results in discards up to 50% of the raw material, consisting of scales, shells, frames, backbones, viscera, head, liver, skin, belly flaps, dark muscle, roe, etc. Besides, fishing operations targeted at popular fish and shellfish species also result in landing of sizeable quantity of by-catch, which are not of commercial value because of their poor consumer appeal. Sensitivity to rapid putrefaction of fishery waste has serious adverse impact on the environment, which needs remedial measures. Secondary processing of the wastes has potential to generate a number of valuable by-products such as proteins, enzymes, carotenoids, fat, and minerals, besides addressing environmental hazards. Fishery wastes constitute good sources of enzymes such as proteases, lipases, chitinase, alkaline phosphatase, transglutaminase,

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2016 Elsevier Inc. All rights reserved.

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hyaluronidase, acetyl glycosaminidase, among others. These enzymes can have diverse applications in the seafood industry, which encompass isolation and modification of proteins and marine oils, production of bioactive peptides, acceleration of traditional fermentation, peeling and deveining of shellfish, scaling of finfish, removal of membranes from fish roe, extraction of flavors, shelf life extension, texture modification, removal of off-odors, and for quality control either directly or as components of biosensors. Enzymes from fish and shellfish from cold habitats are particularly useful since they can function comparatively at lower temperatures thereby saving energy and protecting the food products. Potentials of these applications are briefly discussed.

1. INTRODUCTION Current global production of about 160 million tons of capture and aquacultured fishery products generate significant amounts of processing discards such as scales, shells, frames, backbones, viscera, head, liver, skin, belly flaps, dark muscle, roe, etc. There is an effective need for better management of fishery waste including underutilized fish to address environmental issues and total utilization of the commodity (Arvanitoyannis & Kassaveti, 2008; Rustad, Storrø, & Slizyte, 2011; Venugopal & Shahidi, 1995). Fishery wastes constitute rich sources of proteins including enzymes, polyunsaturated fatty acids (PUFAs), bioactive peptides, carotenoids, minerals, hydroxyapatite, chitin, among others (Sachindra & Mahendrakar, 2015; Venugopal, 2009; Venugopal Menon & Lele, 2015). This chapter will briefly discuss potential uses of enzymes from fishery wastes in seafood processing.

2. ENZYMES FROM FISH PROCESSING WASTE Fishery wastes such as viscera, liver, head, and shell are rich sources of enzymes such as proteases, lipases, transglutaminases (TGases), etc., which have many properties like optimal pH and temperature similar to those of terrestrial organisms (Stefansson, 1998; Trincone, 2011; Venugopal, 2009). Seafood species that survive extremely low temperatures of the polar region possess enzymes having novel features to get adapted to the extreme habitats. These features relate to their isozyme distribution, substrate binding, amino acid sequence, low activation energies, high-specific activities, and thermal sensitivities (Debashish, Malay, Barindra, & Joydeep, 2005;

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49

Huston, 2007; Siddiqui & Cavicchioli, 2006). Some examples are pepsin from gastric mucosa of polar cod having high-specific activity at low temperature and a fish trypsin exhibiting high salt stability (Bougatef et al., 2010; Haard & Simpson, 2006). There is immense scope for exploitation of these enzymes for seafood processing and quality evaluation.

2.1 Fish Proteases Proteases include both endopeptidases and exopeptidases depending on whether they hydrolyze at the interior or side chain of protein molecule, respectively. Proteases from fish and aquatic invertebrates belong to four groups, namely, acidic/aspartic proteases (aspartic acid in the active center, eg, pepsin, cathepsin D), serine proteases (serine in the active site, eg, trypsin, chymotrypsin), thiol or cysteine proteases (calpain, cathepsins B, H, L), and metalloproteases like collagenases having metal in the active site (Shahidi & Kamil, 2001; Sriket, 2014). Trypsins from fish and shellfish have generally high catalytic activities over a wide range of pH and temperature values (Balti, Barki, Bougatef, Khan, & Nasr, 2009; Bougatef, 2013; Bustos, Romo, & Hearly, 1999; Khandagale, Sarojini, Kumari, Suman Joshi, & Nooralabettu, 2015; Khangembam & Chakrabarti, 2015; Kishimura et al., 2010; Klomklao, Benjakul, & Simpson, 2012; Silva et al., 2011; Zamani & Benjakul, 2016). Alkaline proteases from fish viscera exhibit optimum pH of 10 (Sila, Nasri, Bougatef, & Nasri, 2012). Carp and seal chymosin, commonly known as rennin, an acidic protease have high milk clotting activity (Shahidi & Kamil, 2001). Chymotrypsin, an endopeptidase secreted by fish pancreatic tissues has higher specific activity than bovine chymotrypsin (Zhou, Budge, Ghaly, Brooks, & Dave, 2011). Novel acidic proteases from sardine have been characterized (Castillo-Yan˜ez, PachecoAguilar, Garcia-Carren˜o, & Navarrete-Del Toro, 2004; Khaled et al., 2011). Fish lysosomal cathepsins have been isolated from a variety of fishery products (Chere, Delbarre-Ladrat, de Lamballerie-Anton, & Verrez-Bagnis, 2007; Sriket, 2014). Collagenases are present in epithelial, cartilaginous, bony tissues, and digestive tracts of fish and shellfish (Amesen & Gildberg, 2006; Daboor, Budge, Ghaly, Brooks, & Dave, 2012). An acidic serine carboxypeptidase has been isolated from the hepatopancreas of squid (Komai, Kawabata, Tojo, Gocho, & Ichishima, 2007). Table 1 indicates some proteases isolated from different fishery products and their outstanding properties.

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V. Venugopal

Table 1 Proteases from Fish Processing Discards, Their Isolation and Outstanding Properties Some Outstanding Enzyme Fish/Shellfish Properties References

Pepsin

Haard and Simpson Generally low Sardine, capelin, cod, salmon, shark, activation energy, low (2006) and Sriket temperature optimum (2014) mackerel, orange roughy, tuna, trout, carp, harp seals

Gastricin

Hake, Atlantic cod Optimal activity at pH 3

Trypsin

Sardine, capelin, salmon, cod, bluefish, anchovy, Atlantic croaker, carp, aquacultured tilapia, ray fish, cuttlefish, mackerel, threadfin hakeling, red snapper, shrimp, Antarctic krill peel and exudate, smooth hound

Molecular weights about 25 kDa, optimal pH 8–9, and temperature 50–60°C. Threadfin hakeling trypsin has lower optimum temperature and lower thermostability than mammalian trypsin. Smooth hound trypsin has high salt tolerance

Alkaline proteinase

White croaker, chum salmon, tilapia, aquacultured tilapia, viscera of Monterey sardine, goby

Extremely stable in the Sila et al. (2012) pH range of 5–12 and relatively stable toward oxidizing agents

Acidic protease

Sardine, orange roughy

Acidic protease, pH optima 2.5, temperature optima 37–45°C

Castillo-Yan˜ez et al. (2004) and Khaled et al. (2011)

Calpain

Freshwater prawn and fish muscle

Optimal pH 5–7.5, temperature 50°C to
60°C

Chere et al. (2007) and Sriket (2014)

Lysosomal cathepsins type B, H, L

Atlantic white croaker, carp, tilapia, herring, dogfish, rainbow trout, crustaceans, mollusks, squid, prawn muscle

Optimal pH 5–7.5, Catalytic activity possible at low temperatures

Shahidi and Kamil (2001) and Sriket (2014)

Haard and Simpson (2006) Balti et al. (2009), Bougatef (2013), Bustos et al. (1999), Kishimura et al. (2010), Klomklao et al. (2012), Sriket (2014), and Zamani and Benjakul (2016)

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Table 1 Proteases from Fish Processing Discards, Their Isolation and Outstanding Properties—cont'd Some Outstanding Enzyme Fish/Shellfish Properties References

Milk clotting activity Chymotrypsin Carp, capelin, at alkaline pH herring, Atlantic cod, rainbow trout, scallop, spiny dogfish, prawn, sardine Except cathepsin D, an aspartic protease, others are serine or cysteine proteases. Degrade fish myofibrils

Sila et al. (2012) and Sriket (2014)

Aoki, Ahsan Md, Matsuo, Hagiwara, and Watabe (2004), Chere et al. (2007), and Sriket (2014)

Cathepsin D and other lysosomal cathepsins

Capelin, carp, tilapia, chum salmon, squid, white croaker, tilapia, cod, Pacific rock fish

Collagenases

Carp, catfish, cod, Comparable to crab species, pacific mammalian metalloproteinases rock fish, marine bivalve

Amesen and Gildberg (2006), Daboor et al. (2012), and Shahidi and Kamil (2001)

Elastase

Herring, sardine

Causes belly bursting

Felberg et al. (2010)

Chymosin (rennin)

Carp, seal

Optimal pH 2.0–3.5, possesses high milk clotting activity

Shahidi and Kamil (2001)

Alkaline peptidase

Silver mojarra

Optimum activity at pH 8.5, temperature 50 and 55°C

Silva et al. (2011)

2.1.1 Functional Roles of Proteases in Postmortem Fish Muscle Muscle proteases cause degradation of myofibrillar proteins and collagen, causing texture loss, gaping of fish fillets, mushiness in crustaceans, flavor changes, and belly bursting of pelagic fish (Castillo-Yan˜ez et al., 2004; Felberg et al., 2010; Klomklao et al., 2012; Sriket, 2014; Toldra, 2007; Yang, Rustad, Xu, Jiang, & Xia, 2015). A serine protease involved in the activation of polyphenol oxidase (PPO) zymogen, is responsible for blackspot formation in crustaceans (Martı´nez-Alvarez, Montero, & GomezGuillen, 2008). Proteases, adhering in surimi, washed fish meat, causes

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softening, a phenomenon known as “modori.” Applications of proteases in seafood processing will be discussed later.

2.2 Lipases Lipases are a broad family of enzymes that catalyze the hydrolysis of ester bonds in substrates such as triglycerides, phospholipids, cholesteryl esters, and vitamin esters. Lipases exhibit specificity in terms of fatty acids, nature of the alcohol, and stereospecificity. Occurrence, isolation, properties, and physiological role of lipases in aquatic organisms have been discussed (Kurtovic, Marshall, Zhao, & Simpson, 2009; Shahidi & Kamil, 2001). Important applications of lipases in seafood processing include modification of flavor and preferential hydrolysis of ethyl esters of PUFAs (Kamal, Barrow, & Rao, 2015; Kapoor & Gupta, 2012; Toldra, 2007; Venugopal, 2009).

2.3 Transglutaminases TGases are involved in the regulation of cellular growth, differentiation, blood clotting, epidermal keratinization, and stiffening of the erythrocyte membrane. TGase catalyzes acyl-transfer reaction of the γ-carboxamide group of peptide-bound glutamine residues with the ε-amino group of lysine residues and other amines in proteins. When the lysine ε-amino group acts as the acceptor, (γ-glutamyl)-lysine cross-linking occurs between the proteins. TGases have been isolated from several fish species (Binsi & Shamasundar, 2012; Motoki & Seguro, 1998; Shahidi & Kamil, 2001).

2.4 Carbohydrases Carbohydrases isolated from fish and shellfish include chitinases (Proespraiwong, Tassanakajon, & Rimphanitchayakit, 2010), amylase from abalone (Hsieh, Yen, & Jiang, 2008), β-1,3-glucanase from sea cucumber (Zhu et al., 2008), and β-galactosidase from tilapia (Taniguchi & Takano, 2004). Chitinases can be endo- or exochitinases. Endochitinases randomly hydrolyze the internal β-1,4-N-acetyl-D-glucosaminide linkages of chitin, the major crustacean polysaccharide, generating soluble oligomers of GlcNAc. Of the exochitinases, chitobiosidases cleave the dimer, GlcNAc2 and higher chitin polymers, including GlcNAc3 and GlcNAc4, to generate monomers of GlcNAc, while N-acetyl glucosaminidases hydrolyze the terminal, nonreducing N-acetyl glucosamine residues of chitin. The

Enzymes from Fishery Waste for Seafood Processing

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antibacterial enzyme, lysozyme, which recognizes the GlcNAc3–5 for hydrolysis is present in the gastrointestinal tracts and pyloric caeca of shellfish and finfish (Dahiya, Tewari, & Hoondal, 2006; Myrnes & Johansen, 1994).

2.5 Other Enzymes Hyaluronidase, a tissue constituent in animals, which degrades hyaluronate to increase tissue permeability, has been isolated from shrimp shell, Norway lobster, and stonefish found in shallow tropical waters. Enzymes, which have applications in seafood quality evaluation include alkaline phosphatase, urease, 50 -nucleotidase, nucleoside phosphorylase, xanthine oxidase, trimethylamine oxide (TMAO) reductase, and TMAO-demethylase (Benjakul, Klomklao, & Simpson, 2009; Klomklao et al., 2012; Zhu et al., 2009).

2.6 Recovery of Enzymes from Seafood Waste for Seafood Processing Fishery wastes constitute rich sources of diverse proteases and other enzymes such as chitinase, alkaline phosphatase, and hyaluronidase which are abundantly available in the intestines followed by pyloric ceca, pancreatic tissues, hepatopancreas, shell, and other waste components (Sriket, 2014). Being highly perishable, the wastes must be available in good quality in relatively large amounts on a regular basis for recovery of enzymes. Some of the commercially viable isolation processes for enzymes from crude waste extracts include precipitation by salts and polyacrylic acids, isoelectric solubilization/precipitation, ultrafiltration, pH shift, flocculation and membrane filtration, and overcooled acetone extraction (Benjakul et al., 2009; Murado, del Pilar Gonza´lez, & Jose Va´zquez, 2009; Penven, Galvez, & Berge, 2013). Overcooled acetone can recover ray fish collagenases (Murado et al., 2009). Glycation is a novel technique for protein isolation (Sanmartı´n, Arboleya, Villamiel, & Moreno, 2009). Purification strategies of lipases from fishery sources have been reviewed (Kurtovic et al., 2009).

3. APPLICATIONS OF FISHERY ENZYMES IN SEAFOOD PROCESSING Applications of enzymes including fishery enzymes in industrial processing have many advantages over chemical methods such as better process control, environmental and toxicological safety, low energy requirement, and cost. The potential applications of fish enzymes have been reported by several authors (Benjakul et al., 2009; Haard & Simpson,

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2006; Kim & Dewapriya, 2014; Silva et al., 2011; Stefansson, 1998; Venugopal, 2009; Venugopal, Lakshmanan, Doke, & Bongirwar, 2000). Their potential applications for seafood processing are summarized in the subsequent discussion.

3.1 Proteases 3.1.1 Recovery of Proteins from Fish Processing Waste Fishery wastes constitute rich sources of various proteins including myofibrillar and sarcoplasmic proteins and collagen. Collagens from the fins, scales, skins, bones, head, and swim bladders of bighead carp have been extracted using collagenases, pepsin from tuna or trypsin from cod or tuna pyloric caeca (Ahmad & Benjakul, 2010; Amesen & Gildberg, 2006; Jeya Shakila & Jeyasekaran, 2015; Nalinanon, Benjakul, Visessanguan, & Kishimura, 2008; Tschersich & Choudhury, 1998). Pepsin-solubilized collagen (PSC) could be plausible substitutes to mammalian collagen. PSC and its hydrolyzed product, gelatin, have relevant applications in the food, pharmaceutical, and photographic industries (Guillen Gomez, Gimenez, Lo´pezCaballero, & Montero, 2011). 3.1.2 Fish Protein Hydrolyzates Fish protein hydrolysate (FPH) is a breakdown product of fish proteins containing smaller peptides and amino acids. FPH is obtained by treatment of fish meat with trypsin, alcalase, chymotrypsin, pepsin, or other enzymes under controlled conditions of pH and temperatures. Most FPHs are amorphous powders, hygroscopic in nature, containing 81–93% protein, less than 5% fat and 3–8% ash and 1–8% moisture. Lean fish species or their processing wastes are ideal raw material for FPH, which can be used as food binders, emulsifiers, gelling agents, and nutritional supplements. Besides, FPH can function as cryoprotectant and nutritional additive in liquid fertilizer and aquafeed (Chalamaiah, Dinesh Kumar, Hemalatha, & Jyothimay, 2012; Kristiansson & Rasco, 2000; Venugopal, 2009). Seafood proteases from Atlantic salmon and trypsin from fish pyloric ceca have been used for FPH (Kristiansson & Rasco, 2000; Zamani & Benjakul, 2016). 3.1.3 Debittering of FPHs FPHs are often bitter in taste due to the presence of certain peptides. An acidic serine carboxypeptidase isolated from the hepatopancreas of squid can eliminate the bitterness by removing specific amino acids in the carboxy terminal (Komai et al., 2007).

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3.1.4 Bioactive Peptides FPH is a good source of bioactive peptides, which exhibits diverse activities such as antioxidant, antimicrobial, antihypertensive, antiamnesiac, mineral binding, immunomodulatory, and antithrombotic functions (Hayes & Flower, 2014; Kim & Mendis, 2006; Ktari et al., 2013; Vercruysse, van Camp, & Smagghe, 2005; Zhang et al., 2015). Crude extracts of cuttlefish and sardine were used to prepare antioxidant peptides from cuttlefish meat (Ktari et al., 2013). Similarly, fish intestinal protease yielded a phosphopeptide with calcium-binding activity upon treatment of fish bone (Jung, Park, Byun, Moon, & Kim, 2005). Hyaluronidase and elastase inhibitory peptides were prepared from clam (Sutthiwanjampa & Kim, 2015). 3.1.5 Fish Sauce Fish sauce is a popular condiment due to its characteristic flavor and taste, prepared by autolysis of fish by in situ proteolytic enzymes. Its production process can be accelerated by exogenous proteases (Lopetcharat, Choi, Park, & Daeschel, 2001). Addition of squid hepatopancrease accelerated sauce production from capelin (Helgi, Hjalmarsson, Park, & Kristbergsson, 2007). 3.1.6 Ripening of Fermented Fishery Products Ripening contributes to the development of characteristic flavor and soft texture in fermented fishery products during storage, which may be accelerated by exogenous proteases (Bekhit, 2011). 3.1.7 Meat Tenderization Postmortem toughening of meat is a problem that affects its consumer acceptability. Aoki et al. (2004) observed that shrimp protease can tenderize beef. The hard texture of squid rings can be softened by protease treatment, after removal of the tough outer proteoglycans layer of the rings (Melendo, Beltran, & Pedro, 1997). 3.1.8 Control of Curd Formation in Canned Fish Curd formation in canned salmon can be controlled by protease treatment (Gildberg, 1993). 3.1.9 Scaling of Fish Scales can account up to 5% of whole finfish weight. Manual or mechanical scrubbing to remove scales can damage fish having large scales such as haddock, salmon, perch, ocean bream, and silver carp. On the other hand,

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holding the fish in slightly acidified aqueous solution of suitable protease followed by washing helps easy removal of scales. Collagenase from crab hepatopancreas has been used for skinning of squid (Svenning, 1993). 3.1.10 Caviar Production Caviar is cured fish eggs of fish such as white sturgeon, salmon, trout, etc. While manual removal of roe sacks of eggs to prepare caviar is cumbersome, proteases including collagenases can assist for the easy removal of the supportive tissue (Gildberg, 1993; Svenning, 1993). 3.1.11 Extraction of Chitin Shrimp shell discards are an excellent source of chitin, the major component of crustacean and cephalopods shells. Chitosan, obtained by partial deacetylation of chitin is a valuable polysaccharide for its antimicrobial, antitumor, hypocholesterolemic, and immunostimulation, and other functions (Venugopal, 2009, 2011). Traditionally, chitin is extracted by demineralization and deproteinization using strong acids or bases. Alkaline proteases from fish viscera facilitate efficient removal of proteins under mild conditions (Alishahi & Aider, 2011; Sila et al., 2012; Thiago, Cahu´, Santos, Mendes, & Cordula, 2012; Va´zquez et al., 2013). Chitosan because of its antimicrobial and antioxidant activities can be a stabilizer to seafood-based products (Alishahi & Aider, 2011). 3.1.12 Isolation of Carotenoids Shrimp and crab shell waste contain about 150 mg of carotenoids per kg on dry weight basis. The major carotenoid, astaxanthin, from shrimp waste can be recovered after trypsin hydrolysis for use as an ingredient in aquafeed as well as for improving color of salmonid and crustacean species (Alishahi & Aider, 2011; Thiago et al., 2012). 3.1.13 Chondroitin Sulfate and Hyaluronic Acid Chondroitin sulfate (CS) and hyaluronic acid (HA) are glycosaminoglycans having wide biomedical applications. Elasmobranchs fins are the common source of CS. Skate pancreas proteases can be used to isolate CS from skate cartilage. Protease treatment has also been employed for isolation of HA from fish eyeballs (Va´zquez et al., 2013).

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3.1.14 Production of Seafood Flavorings Seafood flavors are in demand to enhance appeal of surimi-based seafood items such as artificial crab meat and fish sausage. Proteases can aid extraction of flavor compounds from crustacean shells and other materials (He, Chen, Li, Zhang, & Gao, 2004; Imm & Lee, 1999). FPH can also be used as model flavor systems by incorporating fish oil and other ingredients (Peinado, Koutsidis, & Ames, 2016). 3.1.15 Viscosity Reduction of Stick Water Stick water is a by-product obtained during the production of fishmeal. During evaporative concentration of stick water, as the dissolved protein increases beyond 25%, increase in viscosity leads to clogging of the evaporator and reduces evaporation. The problem can be addressed by protease treatment (Venugopal et al., 2000). 3.1.16 Control of Enzymatic Browning Enzymatic browning remains a problem for the shellfish, fruit, and vegetable industry Proteolytic enzymes from stomach-less marine species such as crayfish have been recognized to inactivate PPO responsible for blackening and therefore could be an effective alternative to sulfites (Benjakul et al., 2009). Hypotaurine and other sulfinic acid analogs from blue mussel also inhibit PPO (Schulbach et al., 2013). 3.1.17 Production of Aquafeed Feed ingredients containing nutritionally important docosahexaenoic acid (DHA) and eicosapentaenoic acid (EPA) were made by enzymatic digestion of eye or brain tissues from dusky rockfish and salmon (Reppond, de Oliveira, & Bechtel, 2009). Potential uses of fishery waste proteases for seafood processing are summarized in Table 2.

3.2 Transglutaminase 3.2.1 Texture Modification Texture of processed fishery products has strong influence on their consumer acceptance. While additives such as starch and other carbohydrates are conventionally used for texture modification, use of TGase helps texture modification through cross-linking of proteins. TGase-induced setting of surimi has also been suggested as a method for manufacture of restructured

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Table 2 Applications of Proteases for Seafood Processing Processes Benefits References

Protein recovery from fileting waste, mollusk shells, frames of fish as Pollock, etc.

Waste utilization, cleaner Amesen and Gildberg fish frames, and shells as (2006), Jeya Shakila and Jeyasekaran (2015), and calcium source Tschersich and Choudhury (1998)

Extraction of collagen and Lower cost, food gelatin ingredients

Ahmad and Benjakul (2010), Amesen and Gildberg (2006), Nalinanon et al. (2008), and Tschersich and Choudhury (1998)

Extraction of chondroitin Lower cost, important sulfate and hyaluronic acid bioactive compounds

Va´zquez et al. (2013)

Nutritional additives, Preparation of hydrolyzates from fish and flavor enhancers, use in aquafeed shellfish

Benjakul et al. (2009), Chalamaiah et al. (2012), Kristiansson and Rasco (2000), Stefansson (1998), Venugopal (2009), and Venugopal Menon and Lele (2015)

Debittering of protein hydrolyzates

Komai et al. (2007) and Stefansson (1998)

Better acceptance

Preservation of seafood at Chalamaiah et al. (2012), Preparation of Hayes and Flower (2014), antioxidant, antimicrobial chilled temperatures Kim and Mendis (2006), peptides Kristiansson and Rasco (2000), and Ktari et al. 2013 Removal of fish roe sacks Convenient compared to Gildberg (1993) for caviar production cumbersome manual process Ripening of fermented roe

Less time, better yield

Bekhit (2011)

Tenderization of squid and other meat products

Better consumer acceptability

Aoki et al. (2004) and Melendo et al. (1997)

Fish feed digestion

Enhanced absorption

Reppond et al. (2009)

Control of blackening in shellfish

Better consumer value

Benjakul et al. (2009)

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Table 2 Applications of Proteases for Seafood Processing—cont'd Processes Benefits References

Peeling and deveining of shrimp

Simplification of traditional hand peeling

Husain (2010) and Venugopal (2006, 2009)

Membrane removal from Cleaner liver for oil cod liver extraction

Gildberg (1993)

Fish sauce production

Improved yield, flavor, lower fermentation time

Gildberg (1993) and Helgi et al. (2007)

Control of “curd” formation in canned salmon

Better product appearance Gildberg (1993)

Seafood flavor extraction

Recovery of fish flavor from shrimp shell, low cost fish

Toldra (2007), He et al. (2004), Imm and Lee (1999), and Peinado et al. (2016)

Recovery of chitin from shellfish waste for the preparation of chitosan (deacetylated chitin) and chitosan oligosaccharides

Improved, less costly process, energy savings. Chitosan and chitosan oligosaccharides can stabilize seafood-based products. Composite film of chitosan-squid collagen is a good biopackaging

Alishahi and Aider (2011), Kim and Rajapakse (2005), Lin, Lin, and Chen (2009), Thiago et al. (2012), Va´zquez et al. (2013), and Venugopal (2009, 2011)

Recovery of proteins and Improved, less costly carotenoids from shellfish process, energy savings waste

Thiago et al. (2012)

Reduction of viscosity of Better water evaporation, Venugopal et al. (2000) fish meal stick water cost reduction Ripening of salted fish

Better flavor

Gildberg (1993) and Toldra (2007)

Scaling of fish

Efficient process

Stefansson (1998), Svenning (1993), and Tschersich and Choudhury (1998)

Control of enzymatic browning

Inactivates polyphenol oxidase, better color

Benjakul et al. (2009) and Schulbach et al. (2013)

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Table 3 Some Potential Uses of Transglutaminases in Fish Processing

Fish meat paste formulations Fish meat sheet formation Freeze-texturization of fishery products Improvement of binding efficiency of ingredients such as soy proteins Microencapsulation of fish oils in fish proteins Development of protein-polysaccharide coatings and films Raw and processed fish egg products Reduction of drip in thawed frozen fish Surimi and surimi-based fish analogs

low salt fishery products (Motoki & Seguro, 1998). Table 3 gives potential uses of TGases in seafood processing.

3.3 Lipases 3.3.1 Production of PUFAs Specific nutritional benefits of EPA and DHA have promoted research seeking novel lipid sources (Venugopal, 2009). Lipases can be used for enrichment of EPA and DHA of fish oils. The technique involves acidolysis, ie, interesterification where fatty acids of a triglyceride or mixed triglycerides are exchanged or transesterification, ie, exchange of fatty acids of other triglycerides or monoesters (Klinkesorn, Kittikun, Chinachoti, & Sophanodora, 2004; McNeill, Ackman, & Moore, 1996). 3.3.2 Flavor Enhancement Lipases together with muscle cathepsins, calpains, peptidases, and aminopeptidases influence release of flavor compounds (Bekhit, 2011; Toldra, 2007).

3.4 Carbohydrases 3.4.1 Deacetylation of Chitin to Prepare Chitosan Conventional deacetylation of chitin to prepare chitosan by concentrated sodium hydroxide at high temperatures is cumbersome (Venugopal, 2009, 2011). On the other hand, deacetylation by chitinase is less expensive and environmental friendly (Kim & Rajapakse, 2005). Low molecular

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weight chitosan products have also been prepared using chitinase, lysozyme, and cellulase (Lin et al., 2009). 3.4.2 Shelf Life Extension of Fishery Products Dipping fresh shrimp in lysozyme at concentration up to 150 μg/mL retards the growth of spoilage microorganisms due to the action of the enzyme on the mucopeptide structure of microbial cell walls. The enzyme from clam and Arctic scallop exhibited significant antibacterial activity even at lower temperatures. Combination of lysozyme, glucose oxidase, and catalase can enhance fish quality (Myrnes & Johansen, 1994; Ramesh & Lewis, 1980). Application of antibacterial and antioxidant peptides from fishery wastes, as discussed earlier, is an upcoming area for seafood preservation. 3.4.3 Peeling and Deveining of Shrimp A process for loosening the shell of shrimp and removal of visceral mass in clam involves immersion of headless shrimp in a circulating aqueous solution of amylase and sodium bicarbonate followed by washing the shellfish water jets and air to remove shell and the vein (Scott, 1975). 3.4.4 Retention of Color of Cooked and Frozen Shrimp The characteristic yellow color of precooked frozen shrimp or crab as a result of oxidation can be prevented by dipping the cooked shrimp in glucose oxidase. The surface pH is lowered thanks to the generated gluconic acid retarding bacterial spoilage (Scott, 1975). 3.4.5 Other Applications β-Galactosidase, perhaps, is the widely used carbohydrase in food industry to improve sweetness, solubility, flavor, and digestibility (Husain, 2010). Incorporation of carbohydrases in aquafeed can mitigate the adverse effects of nonstarch polysaccharides of the feed (Sinha, Kumar, Makkar, De-Boeck, & Becker, 2011). Glutathione peroxidase can be used to reduce oxidative deteriorative reactions, a major reason for quality loss of flesh foods (Benjakul et al., 2009).

3.5 Quality Control Enzymes such as 50 -nucleotidase, nucleoside phosphorylase, xanthine oxidase, and TMAO reductase are used directly or as component of biosensors for detection of metabolites, toxins, and other contaminants of seafood. Biosensors are analytical devices composed of a biological recognition element

Table 4 Enzymes Other than Proteases and Transglutaminases from Fishery Sources and Their Applications in Seafood Enzymes Source Applications References

Lipases

Atlantic cod, seal, salmon, sardine, Indian mackerel, red sea bream, and others

Shahidi and Kamil Production of omega-3-enriched (2001) and Toldra (2007) triglycerides, improvement of flavor

Chitinases

Shellfish, squid liver, octopus saliva

Alternative to mineral acids to deacetylase chitin

Kim and Rajapakse (2005), Lin et al. (2009), and Proespraiwong et al. (2010)

Amylase

Abalone

Peeling and deveining of shrimp

Hsieh et al. (2008) and Scott (1975)

β-1,3-Glucanase, β-galactosidase

Abalone, scallop, tilapia, sea cucumber

Removal of antinutritive effects of nonstarch polysaccharides in aquafeed

Debashish et al. (2005), Husain (2010), Taniguchi and Takano (2004), and Zhu et al. (2008)

Lysozyme

Arctic scallop shell, Bacteriostatic crab shell agent. Can produce low molecular weight antibacterial chitosans in coordination with cellulase and chitinase

Lin et al. (2009), Myrnes and Johansen (1994), and Ramesh and Lewis (1980)

Catalase, glutathione peroxidase

Marine mussel and Antioxidants. other organisms Combination of lysozyme with glucose oxidase/ catalase helps maintenance of fish quality

Ashie (2012), Debashish et al. (2005), Myrnes and Johansen (1994), and Ramesh and Lewis (1980)

Other enzymes Various fishery (50 -nucleotidase, sources nucleoside phosphorylase, and xanthine oxidase)

Direct measurement of amines, nucleotides, etc., as quality indices or as components of biosensors

Ashie (2012), Venugopal (2006), and Venugopal et al. (2000)

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such as an enzyme, antibody, receptor, or microbe coupled to a chemical or physical transducer. In fishery products, biosensors have potential for proximate analysis, nutritional labeling, determination of pesticide residues, microbial contamination, pathogens, their toxins, among others (Ashie, 2012; Venugopal, 2006; Venugopal et al., 2000). Table 4 summarizes major enzymes (other than proteases and TGases) from fishery source and their applications.

3.6 Uses of Cold-Adapted Enzymes Cold-active proteases facilitate seafood processing at low temperatures such as caviar production (by cold-active fish pepsins or crab hepatopancreas), meat tenderization, and extraction of carotenoprotein (Gildberg, 1993; Haard & Simpson, 2006). Flavor can be improved by spraying cold-adapted proteases onto the surfaces of marine fish and shrimp meat followed by chilled storage improved taste (Alishahi & Aider, 2011). Cold-active lipases facilitate development of various flavors owing to their activities at low temperatures (Birschbach, Fish, Henderson, & Willrett, 2004). TGase from polar fishes can be used for texture modification at low temperatures (Bougatef et al., 2010; Sriket, 2014), while antifreeze proteins from these fishes can inhibit crystal formation during frozen storage of muscle foods (Haard & Simpson, 2006; Venugopal, 2009).

4. COMMERCIAL ASPECTS Enzymes, dominated by carbohydrases followed by proteases and lipases, form a sizeable part of global food and beverage additive market, likely to reach US $2300 million by 2018 (http://www.marketsandmarkets. com). Fish trypsin, chymotrypsin, and cold-active chlamysin (lysozyme) from a marine bivalve are available commercially (Shahidi & Kamil, 2001). Other commercial products include cold tolerant protease from North Atlantic cod, designated as Penzim (http://www.andra.is/), Neptune Aquatein, krill extract (http://ingredientsnetwork.com/ neptune-technologies-bioressources-comp249137.html), heat labile shrimp alkaline phosphatase, cod uracil-DNA glycosylase from Atlantic cod (http:// arcticzymes.com), oyster peptide and protein blends from CIFT, Kochi, India (www.cift.res.in), and other seafood protein extracts (http:// novozymes.com). Potential also exists for protein engineering aimed at for introducing novel and precise sequence specificity into marine enzymes that can result in interesting applications (Pogson, Georgiou, & Iverson,

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2009). Pepsin-extracted and acid soluble collagens and gelatins from fish such as cod bone gelatin have received attention as food ingredients (Guillen Gomez et al., 2011; Karim & Bhat, 2009). In spite of its health benefits, FPHs are not available in good quantities (He, Franco, & Zhang, 2013). Calcitonin, perhaps, is the only bioactive peptide hormone commercially produced from salmon for the treatment of osteoporosis. Enzymatic fish processing has been in vogue in countries like Norway (Raa, 1990). Nevertheless, globally, commercial plants for isolation of fishery enzymes and other value-added products are sparse, in spite of availability of huge quantity of discards (Penven et al., 2013). As pointed out by Olsen, Toppe, and Karunasagar (2014), reasons for low market penetration of fishery by-products including enzymes are nonavailability of high quality waste on a sustainable basis for secondary processing and costs of isolation of ingredients. Nevertheless, with the discovery of new functions and applications, as discussed in this chapter, potential exists to make fishery waste as a source of enzymes to initiate biotechnological approach for seafood processing.

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CHAPTER FOUR

Marine Fungal and Bacterial Isolates for Lipase Production: A Comparative Study H.S. Patnala*, U. Kabilan†, L. Gopalakrishnan{, R.M.D. Rao*, D.S. Kumar*,1 *Indian Institute of Technology, Hyderabad, Telangana, India † School of Bioengineering, SRM University, Kattankulattur, Tamil Nadu, India { PES University, Bangalore, India 1 Corresponding author: e-mail address: [email protected]

Contents 1. Importance of Marine Microbes for the Production of Enzymes 2. Lipase 2.1 History of Lipase 2.2 Lipase Chemistry and Mechanism 2.3 Uses of Lipase 3. Importance of Lipase in Food and Nutrition 3.1 Fat and Oil Industry 3.2 Dairy Industry 3.3 Lipases as Biosensors for Food Industry 4. Commercial Applications of Lipase 5. Production of Lipase by Microbes: Importance of Marine Microbes 6. Production of Lipase by Marine Bacteria 7. Production of Lipase by Marine Fungi 8. Methods for Production of Lipase 8.1 Solid-State Fermentation 8.2 Submerged Fermentation 9. Conclusion Acknowledgments References

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Abstract Lipases, belonging to the class of enzymes called hydrolases, can catalyze triglycerides to fatty acids and glycerol. They are produced by microbes of plant and animal origin, and also by marine organisms. As marine microorganisms thrive in extreme conditions, lipases isolated from their origin possess characteristics of extremozymes, retain its activity in extreme conditions and can catalyze few chemical reactions which are impossible otherwise relative to the lipase produced from terrestrial microorganisms. Lipases are useful Advances in Food and Nutrition Research, Volume 78 ISSN 1043-4526 http://dx.doi.org/10.1016/bs.afnr.2016.06.001

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in many industries like detergent, food, leather, pharmaceutical, diary, etc. Few commercial enzymes have been developed and the use of them in certain industries like dairy, soaps are proved to be beneficial. There are few research papers reporting the production of lipase from marine bacteria and fungi. Lipase production involves two types of fermentation processes—solid-state fermentation (SSF) and submerged fermentation (SmF). Although SmF process is used conventionally, SSF process produces lipase in higher amounts. The production is also influenced by the composition of the medium, physiochemical parameters like temperature, pH, carbon, and nitrogen sources.

1. IMPORTANCE OF MARINE MICROBES FOR THE PRODUCTION OF ENZYMES The mineralization of complex organic matter and contribution to the secondary production in the sea is carried out by the marine organism which takes an active part in the marine environments. These microorganisms help to degrade the many materials other than the dead plants and animals which are present in this marine environment; these include complex polysaccharides such as lignin, cellulose, pectin, xylan, starch, proteins, fats, sugar, urea, aromatic and aliphatic hydrocarbons, and several other organic compounds (Chandrasekaran & Kumar, 2010). Due to habitual properties, the marine microorganisms become active in the extreme conditions prevalent in the marine environments. These organisms had to face extreme environmental challenges and have adapted to survive as the course of evolution because of which they produce unique metabolites which are seldom reported from terrestrial sources. These extreme conditions include stability and activity at high temperature or pH, high tolerance to saline conditions, active in the presence of different chemicals or even organic solvents. It is also reported that marine enzymes have properties that help it to catalyze certain chemical reactions which are considered to be impossible using other catalysts. General Biotechnological perspective has recognized these characteristics that are shown by these marine microorganisms which arise out of its habitual characteristics, it has been observed that blending of any two such features into a single enzyme is extremely useful in the development of unique bioprocesses. For example, thermostable alkaliphilic proteases are used as additive in laundry detergents and these enzymes can be useful for dry cleaning if they can work in the presence of organic solvents. Another example is in the production of biofuel from seaweed biomass if the enzyme is saline environment active. It helps as the enzyme will be solvent tolerant and also thermally stable (De Romo & Borgstein, 1999).

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2. LIPASE Lipase is an enzyme that catalyzes the hydrolysis of triacylglycerol to yield diacylglycerol and a fatty acid anion. It belongs to the family of hydrolases that catalyze the breakdown of fats. This enzyme is involved in the digestion of fats in our bodies. It is present in glands of the mouth and the pancreas. The enzyme is also detected in the gastric juices and lysozymes. There are two types of lipases: acidic and alkaline lipases. The acidic lipases consist of lingual lipase and gastric lipase, while the alkaline lipases consist of pancreatic lipase. Lingual lipase is said to be the first step in the digestion of lipids. It mostly works in the case of infants as the pancreatic enzymes would not be fully developed.

2.1 History of Lipase Claude Bernard, father of modern physiology, made the discovery of pancreatic lipase in 1848. He isolated pancreatic juice from a dog after 22 failures (Davenport, 2013). He then took tallow from a candle on his work table and added it to the pancreatic secretion. He noticed the emulsification of the fat from the candle, thus discovering the presence of lipase in the pancreas. Today, it is well known that pancreatic lipase aids in the absorption and digestion of fats in the human body. It was determined at the beginning of the 20th century that glycerol extracts of the gastric mucosa contained a lipase compound that attacked emulsions of egg yolk, milk, cream, olive oil, and pig fat. It was only in 1973 when Margit Hamosh and Robert Scow brought Gastric lipolysis to the foreground. When they were working, they found that the serous glands of the rat’s tongue contain and secrete a lipase that is active against long-chain triglycerides which are present in the soft palate, in the anterior and pharyngeal tongue, and in lateral and pharyngeal glands (Ray, 2012).

2.2 Lipase Chemistry and Mechanism Lipase breaks down the triglycerides to diglycerides, diglycerides to monoglycerides, and then monoglycerides to fatty acids and glycerol (Figs. 1 and 2). Generally, a lipase exists in two forms: active and inactive form. This characteristic of the enzyme is controlled by a lid-like structure that covers the catalytic site of the enzyme (Fig. 3). The active enzyme has its lid open and allows the substrate interaction in the binding site for catalysis. Catalytic

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H.S. Patnala et al.

OH2

Lipase

+ O C R1

O

R2

R2 H

OH

O H N

H R

–

H

O H Nucleophile

C O

O R+

R1

Active serine Lipase O C R1

Nucleophile

Fig. 1 Bioaction of lipase with nucleophilic targeted carbonyl function (Kumar & Ray, 2014).

domain of lipase contains nucleophile serine residue which facilitates this reaction. Closed lid denotes that this residue could not attack the substrate for the substrate to access the catalytic pocket; a water-lipid surface is required to modulate the lid which helps to open the lid thereby exposing the catalytic site (Ali et al., 2013).

2.3 Uses of Lipase There are a large number of biocatalysts that are being used across the field of biotechnology, lipases being one of them, lipases are used in dairy, food, detergents, textile and pharmaceutical, cosmetic and biodiesel industries. The basic function of lipases is the removal of fatty acids and so is used as a household detergent and in an industrial laundry and for cleaning clogged drains.

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Marine Fungal and Bacterial Isolates for Lipase Production

O CH2

OH

C O

CH2

R1

CH2

O

C O

CH2

R2

CH2

O

C

CH2

R3

+ H2O

Triacyl glycerol CH2

Action by lipase

OH O

CH2

O

C O

CH2

R2

CH2

O

C

CH2

R3

O + H2O +

OH

CH2

O

C

CH2

R1

Free fatty acid

Diacyl glycerol CH2 OH CH2

HO

Action by lipase O

O C

+ H2O + CH2

R3

HO

C

CH2

R2

Free fatty acid

Monoacyl glycerol Action by lipase CH2

OH

CH2

OH

O

CH2

OH

+

HO

C

CH2

R3

Free fatty acid

Glycerol

Fig. 2 Conversion of triglycerides to glycerol and free fatty acids by lipase (Kumar & Ray, 2014).

Protein lid open

Substrate

Enzyme substrate interaction

Product

No product

Protein lid closed

Substrate

No interaction

Fig. 3 Pictorial representation of action of lid with respect to enzyme activity.

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3. IMPORTANCE OF LIPASE IN FOOD AND NUTRITION 3.1 Fat and Oil Industry Foods consist of fats and oils, one of the properties of lipases include the ability to modify properties of lipids which is accomplished by altering the fatty acid chains in glycerides, the alteration may be a change in location or complete replacement with a new one, whereby bringing about a change in its constituents to produce a much higher valued compound (Salihu & Alam, 2012). The common use of commercial lipases includes flavor development in dairy products and processing of meat, fruits, vegetables, etc. Phospholipases are used in the production of mayonnaise from egg yolk; the treatment of the egg yolk increases the emulsifying capacity and heat stability and can be used in the processing of custard, baby foods, and in dough preparation. It is also used in lecithin modification and for oil degumming of vegetable oils, processing of sauces like hollandaise, bearnaise, and cafe de Paris.

3.2 Dairy Industry The primary use of lipases in the dairy industry is to enhance the flavors of various cheeses by the modification of chain lengths of the fatty acids, this kind of enzyme-modified cheese are used to produce a much-concentrated flavor in the presence of enzymes at higher temperatures. The concentrated flavor contains as much as 10 times higher fat as compared to that in normal cheese (Ferreira-Dias, Sandoval, Plou, & Valero, 2013). Cheese ripening is a gradual enzymatic reaction in which freshly worked curd is modified to the desired final ripe cheese texture and flavor.

3.3 Lipases as Biosensors for Food Industry Triacylglycerol is blood cholesterol enhancing compound and it is important to in food industries like fats and oils, beverages, pharmaceutical to be able to effectively quantify the presence of this compound and for this case immobilized lipases are considered to be fast, efficient, accurate, and cost effective as a sensor. The lipase used generates glycerol from the

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Marine Fungal and Bacterial Isolates for Lipase Production

Table 1 Application of Lipase in Food Industry Industry Action Mechanism

Cheese industry

To improve flavor To accelerate cheese ripening

References

Kumar and Ray Lipase releases fatty acids from triglycerides that help (2014) and Ray (2012) in flavor enhancement The presence of lipase causes steady increase in the concentration of fatty acid liberation and total soluble nitrogen and aids in ripening El-Hofi, El-Tanboly, and Abd-Rabou (2011) and Ferreira-Dias By converting the positions et al. (2013) of fatty acids present in the lipids, the oils can made into margarine by increasing melting points

Fats and oil industry

To convert cheap oil to nutritive oils To produce lowcalorie oils To convert physical properties of oils and hard butter

Lipases help to modify the properties of lipids by hydrolysis, esterification, and transesterification

Egg industry

For production of mayonnaise, custard, and eggyolk treatments

Phospholipases hydrolyze Kumar and Ray (2014) the egg lecithin and isolecithin in order to increase the emulsifying capacity and heat stability El-Hofi et al. (2011) and Ferreira-Dias et al. (2013)

Nutraceuticals To improve and functional health benefits in food like food antioxidants and other bioactive compounds

Lipase acts as biocatalyst and is immobilized on a membrane in reactor to bring about interesterification. This improves stability of biocatalyst and there will be better control of reaction

Meat industry

Lipase breaks down fat and Ferreira-Dias reduces the calories present et al. (2013) in meat products

To remove excess fat from fish and meat

triacylglycerol present in the sample and to quantify the amount of glycerol released by either the chemical or enzymatic methods (Table 1). The immobilization is done on the pH/oxygen electrodes in combination with glucose oxidase which acts as the biosensor (Ferreira-Dias et al., 2013).

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4. COMMERCIAL APPLICATIONS OF LIPASE Lipases have a wide range of applications on the commercial scale and it is considered as a lucrative business (Nakajima, Snape, Khare, & Gupta, 2000). The usage of lipases varies because of its varied features which include activity under mild conditions, high stability in organic solvents, a broad range of specificity in the case of substrates, and selectivity in the case of catalysts. The use of lipases also extends to other fields like the production of cocoa butter equivalent, “Betapol” a commonly used substitute for the human milk fat, polyunsaturated fatty acids rich/low-calorie lipids which have pharmaceutical applications and in the production of biodiesel from vegetable oils (Jaeger, Dijkstra, & Reetz, 1999; McNeill, Shimizu, & Yamane, 1991). Other applications include the manufacture of soap as seen in Miyoshi Oil & Fat Co., Japan where Candida cylindracea is used for this purpose (Hasan, Shah, & Hameed, 2006). It was observed that the enzymatic process yielded a superior and cheaper product than produced using the conventional method. Lipolase was the first introduced commercial lipase in 1994, it was extracted from a fungus Thermomyces lanuginosus, in later stages two other bacterial lipases were introduced, which were produced by Anna University, Chennai, India and these were named as Lumafast and Lipomax each from Pseudomonas mendocina and Pseudomonas alcaligenes, respectively (Sharma, Chisti, & Banerjee, 2001). Bakery is an industry which requires a wide range of enzymes for the preparation of a variety of products to fulfill the needs of the bakery trade and this is mostly done by Millbo S.p.a (Italy) which has a supply of different enzymes used to replace the emulsifiers used to increase the volume in bread and bakery (Gutierrez, Jose, & Martı´nez, 2009). Unichem International, a Spanish enterprise specializes in the production of enzymes that can be used as an emollient in personal care products like skin creams, bath oils, etc., which basically contains palmitate products. WW07P is a specially formulated product of Oasis Environmental Ltd., which is used in the biological wastewater treatment especially of those waters which contain high content of greases, fats, and oils, another specific property of this compound is the fact that it contains surface tension depressants which loosen and liquefy heavy grease deposits which further helps in biodegradation (Saxena et al., 1999) (Tables 2 and 3).

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Marine Fungal and Bacterial Isolates for Lipase Production

Table 2 Commercial Applications of Lipase in Different Industries Industry Action Mechanism Advantage References

Cleaning As detergents Lipase can industry remove any oil stains and can remain active in harsh conditions

They can remain unscathed against harsh washing conditions and surfactants

Paper and To remove pitch from pulp industry wood

Lipase hydrolysis It helps in easy triglycerides thus paper manufacturing removing the pitch

Biodiesel To produce industry biodiesel

Lipase used as immobilized biocatalysts which help in transesterification

Textile industry

Bioleaching and In the mixture for desizing along Biodesizing are with α-amylase eco-friendly processes

For desizing denim and also for leaching

This helps in reusability of lipases and also reduces the operational temperature

Aravindan, Anbumathi, and Viruthagiri (2007) and Ferreira-Dias et al. (2013) Ferreira-Dias et al. (2013), Nakajima et al. (2000), and Aravindan et al. (2007) Ferreira-Dias et al. (2013) and Jaeger and Reetz (1998)

Jaeger et al. (1999)

Table 3 List of Companies Using Lipase Obtained from Various Species (Hasan et al., 2006) Name of the Product Name/ Company Application Obtained from Species

Candida cylindracea

Miyoshi Oil and Fat Co., Japan

Use lipase in manufacture of soaps

Rakuto Kasei Israel Ltd.

Produces enzymes for textile industry NM

Gateway Pro Clean Inc., USA

Offers laundry programs utilizing enzyme technology

NM

NM

Kinetic resolution of racemic flurbiprofen

Candida antarctica

Enzymatix UK

Uses lipase in biotransformations

NM Continued

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H.S. Patnala et al.

Table 3 List of Companies Using Lipase Obtained from Various Species (Hasan et al., 2006)—cont'd Name of the Product Name/ Company Application Obtained from Species

Candida cylindracea

Croda Universal Ltd.

Uses lipase in making cosmetics

Maps(India) Ltd.

Provides a range of lipases to work in NM different pH conditions

Novozyme, Denmark

Acid bating of fur and wool

NovoCor ABL and NovoCor ADL

Novozyme, Denmark

Transesterification of crude soybean oils for biodiesel production

Novozyme 435

NM, not mentioned.

5. PRODUCTION OF LIPASE BY MICROBES: IMPORTANCE OF MARINE MICROBES Lipase is one of the biomaterials which received wide attention as it carries out biochemical reactions in both aqueous and nonaqueous media. Lipases may be obtained from plant, animal, and microbial sources. However, commercial applications are using microbial lipases because they can be easily cultivated, are relatively stable and can act as a catalyst for a wide range of hydrolytic and synthetic reactions (Saxena et al., 1999). Lipase is an enzyme that can be secreted by a wide range of microorganisms. Fungi and bacteria may secrete lipase to facilitate nutrient absorption from the external medium. The production of lipase requires carbon and nitrogen sources as required by any fermentation process and they play an important role for an efficient production (Rajesh kumar, Mahendran, & Balakrishnan, 2013). The carbon sources widely used are mostly lipid derivatives. Moreover, simple sugars like glucose, sucrose are also used as carbon sources for the lipase production. Nitrogen sources include peptone, yeast extract (Ire, Ezediokpu, & Okerentugba, 2015). The production of lipase is higher when lipid sources are used as carbon sources. Along with carbon sources, the production rate is enhanced when inducers like vegetable oils, tributyrin, hexadecane are used (Zarevu´cka, 2012). The physical parameters like pH, temperature, etc., also influence the production rate. This varies from species to species, for example, for fungal strains like Fusarium sp., the optimum pH and temperature are reported to be 2.5 and 30°C, respectively, while for bacterial species like Bacillus subtilis the optimum pH and temperature are 7.0

Marine Fungal and Bacterial Isolates for Lipase Production

81

and 30°C, respectively (Ire et al., 2015; Pallavi, Chandra, Reddy, & Reddy, 2014). In the same way, marine microorganisms have its own requirements for the production of lipase. They play an active role in degradation pathways of organic matter which is part of their metabolism to produce secondary metabolites in marine environments. Most of the marine microorganisms are extremophiles which live in harsh marine environments like deep ocean hydrothermal vents, polar ocean, extreme saline bodies, etc., such microorganisms produce extremozymes which retain its activity in diverse conditions (Chandrasekaran & Kumar, 2010) (Fig. 4). The production of lipase was first reported in Penicillium oxalicum and Aspergillus flavus in 1935. Later, four cold-adapted lipases were screened from Moraxella sp. which was obtained from Antarctic seawater with optimum growth temperature at 25°C and the lowest secretion temperature reached 3°C. Another group of researchers isolated nine lipase producing strains from 427 strains of yeast in which some are capable of hydrolyzing different oils. A novel extracellular phospholipase C which is highly specific in hydrolyzing phosphatidyl choline was isolated from marine Streptomycete and showed maximum enzyme activity till 40°C and pH 8.0 (Zhang & Kim, 2010). Production of lipase by marine bacteria and fungi in particular are discussed in the following sections.

6. PRODUCTION OF LIPASE BY MARINE BACTERIA Lipases used for commercial and industrial applications are mostly from bacterial sources. They belong to the family of serine hydrolases and contains α/β hydrolase fold in their core structure. They also have a catalytic triad (serine, aspartate/glutamine, and histidine) in its structure. They are more of inducible enzymes which require inducers made of some form of oil, fatty acids, etc., to carry out the production as the activity of the lipase is constricted against water insoluble substrates (Dalmaso, Ferreira, & Vermelho, 2015). Very few lipases which produced constitutively are reported. They are mostly extracellular and secreted out in the culture medium but in some cases intracellular and membrane bound lipases are also reported. The initial stages in production of lipase vary from organism to organism but generally it occurs during late logarithmic or stationary phase of the growth cycle in an organism (Shah & Bhatt, 2011). Bacterial lipases are used in many industries like dairy industries, pharmaceutical industries, laundry industries, etc., because they are easy to cultivate, nontoxic, and ecofriendly in nature (Boonmahome & Mongkol thanaruk, 2013). Most of

Extremophiles

Psychrophiles

Thermophiles

Acidophiles

Alkaliphiles

Halophiles

Barophiles

0–12°C

90–113°C

Low pH: −0.06 to 4

High pH: 8.5 to 12

High salt content: > 15%

High pressure: > 500 atm.

Fig. 4 Types of extremophiles and their characteristics.

Marine Fungal and Bacterial Isolates for Lipase Production

83

the lipase producing bacteria are isolated from terrestrial origin while bacterial strains from marine environment which is subjected to diverse form of flora and fauna are yet to be explored (Doles, Sarita, & Chandrasekaran, 2014). Bacteria from the marine origins are extremophiles which can thrive in extreme temperatures, pH, salinity, etc. It has been reported that bacterial lipase from genus pseudomonas has a unique property of high thermal stability, high enantiomer sensitivity, and retaining its activity at broader range of temperatures and pH. Lipase from Pseudomonas otitidis isolated from marine sediments retained its activity at temperatures ranging from 30°C to 80°C and pH ranging from 5.0 to 9.0. The maximum lipase production of 1980 U/mL was observed at pH and temperature 7.5 and 35°C, respectively. After purification, the productivity of lipase increased to 5647 U/ mL with 8.4-fold of purity (Ramani, Saranya, Jain, & Sekaran, 2013). Another bacterial genus called Thermococcus which has been isolated from hydrothermal vents or freshwater springs thrives in a condition with 1–3% of salt concentration. Thermococcus sibiricus is a hyperthermophile obtained from Samotlor oil reservoir (western Siberia) contains 15 genes encoding for lipase/esterase out of which four enzymes are considered to be extracellular enzymes. Other hyperthermophilic bacteria from which the lipase activity is reported are Archaeoglobus fulgidus, Pyrobaculus calidifontis, and Sulfolobus sp. like S. solfataricus, S. acidocaldarius, and S. shibatae (Renge, Khedkar, & Nandurkar, 2012). A thermostable lipase was isolated from species belonging to genus Salinivibrio which retained its activity for 30 min at 80°C and pH 7.5–8.0. It also tolerated a range of 0.1–3.0 M NaCl concentration (Dalmaso et al., 2015). A cold-active lipase isolated from bacterium Colwellia psychrerythraea showed optimum activity around 25°C and pH 7.0 and its activity was retained at temperatures as low as 7°C (Do et al., 2013). Similarly, another cold-active extracellular lipase isolated from psychotropic Yersinia enterocolitica showed activity at temperature ranging from 0°C to 60°C with optimum activity at 37°C. Ca2+ had regulatory effect on the enzyme (Ji et al., 2014). Other than pH and temperature, the production of lipase in bacteria is influenced by carbon and nitrogen sources, inorganic salts, oxygen, detergents, etc. (Seghal Kiran, Nishanth Lipton, Kennedy, Dobson, & Selvin, 2014). Lipase production was enhanced in Burkholderia cenocepacia ST8 with a medium containing peptone as nitrogen source (Ananthi, Ramasubburayan, Palavesam, & Immanuel, 2014). Since lipase is an inducible enzyme; it is produced only in the presence of lipid sources like fatty

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acids, tweens, hydrolysable esters, triacylglycerols, etc. However, there are other sources of sugar, polysaccharides which also have positive effect on lipase production. Lipase production in thermophilic Bacillus sp. increased in the presence of olive oil as a carbon source. Other papers also stated that glucose in the culture medium inhibits the enzyme production through catabolic repression (Sarethy et al., 2011). Metal ions which are another key factor reported to enhance the production of lipase. The most favorable metal ions are Ca2+ and Mg2+. The lipase activity is also enhanced by Ca2+ and Mg2+ and partially inhibited by Cu2+, Zn2+, Ba2+, Pb2+, Fe2+, and Mn2+ in cold-active lipase isolated from Antarctic sea ice bacteria Pseudoalteromonas sp. NJ 70 (Huang, Locy, & Weete, 2004). In the predicted model of lipAMS8, there are metal ions including six atoms of Ca2+ and one atom of Zn2+. Metal ions may also contribute to substrate activation and electrostatic stabilization of enzymes (Sarethy et al., 2011). On the whole, the production of lipase in the bacteria is influenced by more than one factor.

7. PRODUCTION OF LIPASE BY MARINE FUNGI Marine fungi are also being explored for the presence of extracellular enzymes. The fungi and yeast are ubiquitous like bacteria in the air, water, and soil. Nearly, 10,000 species of fungi are said to be habitating in the deep stretches of oceans, out of which only 600 species of fungi have been reported in the oceans, providing the scope for more research. The fungal microbes are known to be good choices for production of secondary metabolites against their terrestrial counterparts, because they have more resistance to physical conditions. They are generally more tolerant toward temperature, pH, salinity, and other extreme conditions which cannot be associated with terrestrial fungi. Due to the presence of such extreme genes within their genome, the translated products are much more stable toward temperature, pH, pressure, and salinity conditions. Therefore, they can be exploited for biotechnological applications. The enzymes that these organisms produce can be used in various industrial sectors of food, textile, cleaning, environmental, and pharmaceuticals. Several research works are going on in the use of fungi for enzyme production. The source of microbe is first washed with sterile seawater. However, Passarini, Santos, Lima, Berlinck, and Sette (2013) used mercuric chloride as a sterilizing agent on the source and the results were not as good as with plain sterile sea water. This could perhaps be attributed to the fact that mercuric chloride could have killed a few microbes present in the source.

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85

The isolates are grown in the culture media and there are various protocols for the different types of fungi (Bonugli-Santos et al., 2015). There can be different screening procedures to identify or select the target microbe. Generally, colorimetric assays are done for microbial enzymes. The purification processes that are adopted for marine fungi are mostly based on ammonium sulfate precipitation, but even ion exchange chromatography is used. Lipase is a hydrolase that is produced by almost all living organisms as it is the primary fat degradation enzyme. The production of lipase from marine microbes has been vast, but research in the isolation of lipase from marine fungi is still evolving. For instance, Aspergillus awamori isolated from the Arabian Sea waters of the Indian coastline (Basheer et al., 2011) was used through the submerged fermentation (SmF) method with an optimized medium. The optimum media components were found to be soybean 0.77% (w/v), ammonium sulfate (0.1 M), potassium dihydrogen phosphate (0.05 M), rice bran oil 2% (v/v), calcium chloride (0.05 M), PEG 60000.5% (w/v), NaCl 1% (w/v), and an inoculum at 1% (v/v). The media had a pH 3 and was incubated at 35°C for a period of 5 days. Maximal lipase production reached within 36 h while the maximum specific activity of 1164.63 U/mg protein was detected at around 108 h. The lipase was partially purified using ammonium sulfate precipitation and then by ion exchange chromatography. The final yield turned out to be 33.7%. Isolated marine fungi were isolated from the Arabian Sea for extracellular lipase and other enzyme activities (Smitha, Correya, & Philip, 2014). The optimum conditions were reported to be peptone (0.5 g), beef extract (0.3 g), agar (2 g), seawater (100 mL), gelatin at 2%, starch (1%), colloidal chitin (5%), cellulose powder (0.5%), tributyrin (1%) under pH 7.2 for a period of 3–5 days. This research group did not search for specific fungi for lipase production but looked at all the fungi from the marine environment of the Arabian Sea that could produce lipase. They found out that species like Penicillium, Aspergillus, Scopulariopsis, Trichoderma, Endomysis, and others could produce lipase enzyme. Out of a total of 180 cultures, around 60.2% displayed the fat degrading enzyme, among which Penicillium was found to show maximum activity. Marine actinomycetes were screened for lipase and for other enzyme activities using a nutrient broth (8 g/L), sodium chloride (4 g/L), agar (100 g/L), olive oil (31.25 mL), 10 mL phodamine B 0.001 % (w/v) at pH 6 and 60°C (Selvam, 2011). The culture was incubated at 28°C for 72 h and the colonies that fluoresced orange under UV were screened as positive lipase producers. Out of the 56 isolated strains, three different strains showed maximum lipase activities. Actinomycete strain 3, Actinomycete

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strain 10, Actinomycete strain 28 showed 586 U/mL, 560 U/mL, and 700 U/mL, respectively. Marine actinomycetes were isolated from agro-industrial wastes for lipase production by solid-state fermentation (SSF) (Suji & Sivaraj, 2014). Ground nut cake and coconut cake were used along with three different carbon sources in dextrose, fructose, and sucrose. Three different nitrogen sources like urea, ammonium nitrate, and peptone at various pH ranging from 5 to 9 and temperatures of 30°C, 37°C, and 47°C. Around eight actinomycetes were reported to contain the lipase activity and five strains showed the maximum activity. The optimum media contents were found to be ground nut cake, ammonium nitrate, sucrose, pH 6, and a temperature of 30°C. The earlier techniques were based on simple isolation and culture of specific marine fungal organisms for lipase production. More novel methods have been adopted for lipase production from marine fungi. Recombinant technology is the new technology which allows engineering the proteins. It is the technology where any gene can be introduced into another with the help of vectors. Candida antarctica which produces lipase A and lipase B has been widely employed in this field. Eom et al. engineered CalB gene of Candida antarctica onto Pichia pastoris strain X-33 (De Marı´a, Sa´nchezMontero, Alca´ntara, Valero, & Sinisterra, 2005; Eom et al., 2013; Yang, Liu, Dai, & Li, 2013). The Pichia expression plasmid of pPICZαA was used. A lipase activity of 5.4 g/L and 57.9 U/L was observed toward p-nitrophenylpalmitate. Hence, the fungal microbes are being exploited for lipase production. They are mainly used because of their extreme tolerance capacity, ease in production, and good physiological characteristics. The lipase from fungi can be used in a variety of industrial sectors (Table 4).

8. METHODS FOR PRODUCTION OF LIPASE Production of lipase involves fermentation processes of mainly two types: SSF and SmF. Usually for bacteria and yeast, SmF is recommended because bacteria require high moisture content, while fungi can be grown in both SmF and SSF.

8.1 Solid-State Fermentation SSF is a fermentation process in which a solid nonsoluble material is used which acts as a physical support as well as a nutrient source for the growth of microorganisms. The technique involves growth of microorganisms on

Table 4 Examples of Different Types of Extremophilic Bacteria and Fungi, Their Requirements and Characteristics Optimum Parameters Required for Lipase Production

Bacterial Species

Pseudomonas otitidis

pH

Temp. Carbon (°C) Source

7.5 35

Glucose

Nitrogen Source

NM

a

Substrate/Inducers

Sunflower oil waste

Enzyme Activity (U/mL) Metal Ions

1980

Ca

2+

Reducing Agents/ Detergents/Others

References

Triton X-100, octane,

Ramani et al. (2013)

p-Nitrophenylcaprate

Dalmaso et al. (2015)

Photobacteriumlipolyticum 9.0 25

Tryptone Yeast extract Olive oil

NMa

NMa

Oceanobacillus sp. PUMB02

Sucrose

Yeast extract Olive oil

58.84

Ca2+, Fe3+, Mg2+ p-Nitrophenylpalmitate Seghal Kiran et al. (2014)

Maltose

Ammonium Tributyrin hydrogen carbonate

326.03 NM

Burkholderia cepacia S31 9.0 70

NMa

NMa

Emulsified olive oil, glycerol

226.1

Mg2+, Ca2+, Tween 80, Ethyl acetate Sarethy et al. Mn2+, K +, Na + (2011)

Staphylococcus lipolyticus

7.0 10

NMa

NMa

Olive oil

NMa

Ca2+

Glycerin, casein

Arora (2013)

Aeromonas hydrophila

9.0 37

Tween 80

Beef extract Fish waste

11.2

NM

Acetone

Neelambari, Vasanthabharathi, Balasubramanian, and Jayalakshmi (2011)

Bacillus aerius

8.0 55

Cotton seed oil

Yeast extract Cotton seed oil

0.164

Zn2+, Mg2+

Tween 20, p-nitrophenylpalmitate

Saun, Mehta, and Gupta (2014)

8.0 30

Bacillus cereus MSU AS –

37

Gingelly oil

Ananthi et al. (2014)

Continued

Table 4 Examples of Different Types of Extremophilic Bacteria and Fungi, Their Requirements and Characteristics—cont'd Optimum Parameters Required for Lipase Production

Bacterial Species

pH

Temp. Carbon (°C) Source a

Psychrobacter sp. C18

8.0 30

NM

Bacillus smithii BTMS 11

8.0 50

NMa

6.0 20

Glucose

Nitrogen Source a

Substrate/Inducers

Enzyme Activity (U/mL) Metal Ions

Reducing Agents/ Detergents/Others

+

References

p-Nitrophenylpalmitate 623.3

K

p-Nitrophenylmyristate, Chen, Guo, and Tween 80, acetone Dang (2011)

NMa

Sesame oil

360

Co2+

β-Mercaptoethanol

Lailaja and Chandrasekaran (2013)

Urea

Corn oil

68

NMa

Triolein, olive oil

Cho et al. (2007)

NM

Fungal Species

Penicillium chrysogenum

a

a

a

Geotrichum marinum

8.0 40

NM

Aspergillus awamori

7.0 40

NMa

NMa

Soybean meal, rice bran 495 oil

Rhizopus homothallicus

7.5 40

Lactose

Urea

Rhizopus oryzae

5.0 45

Sucrose

Peptone

a

NM, not mentioned.

NM

Yeast extract, olive oil

NM

+

2+

Na +, K , Ca , Mg2+

Taurocholic acid, Huang et al. 3-[(3-cholamidopropyl) (2004) dimethylammonio]1-propanesulfonate

NMa

NMa

Basheer et al. (2011)

Olive oil, sugar cane bagasse

10,700 Mn2+

NMa

Diaz et al. (2006)

Olive oil

NMa

NMa

Salleh et al. (1993)

NMa

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porous particulate media with low moisture content in Petri plates, conical flasks, etc. This property of SSF is best suited for the growth of filamentous fungi than any other microorganisms. In contrast, many studies have also reported successful growth of bacteria in SSF. Certain drawbacks of SSF are the heat and mass transfer rates. The common substrates used for SSF are wheat bran, rice bran, and sugar cane bagasse. Physical and chemical parameters play an important role in any fermentation process. Aeration in the chamber, moisture content, pH, and temperature of the substrate in SSF are affected by the growth of microorganisms and heat transfer capacity is very low but this property of SSF can be exploited for the production of thermophilic microorganisms (Mienda, Idi, & Umar, 2011; Raimbault, 1998). To overcome these, new strategies are designed like air pressure pulsation and agitation for heterogeneous substrate utilization and product accumulation. In statistical comparison of lipase production using both, SSF and SmF, SSF showed better yield compared to SmF. The physiochemical parameters and medium optimizations are done using various statistical techniques (Passarini et al., 2013). Microorganisms require an effective medium for its growth that can be optimized through recent developed techniques like Plackett–Burman design (PBD), response surface methodology (RSM), and one factor at time, etc. These techniques help in increasing the productivity of the any bioprocess techniques, interaction and the effect of various physiochemical characteristics can be determined using minimum number of experiments. Optimization of the medium for the production of lipase is achieved through sequential steps of screening using PBD followed by optimizing the important components and verifying the developed medium using RSM (Nakajima et al., 2000). Using these optimization methods, lipase production in pseudomonas otitidis, a marine strain showed optimal lipase activity of 1980 U/mL with optimum conditions (Basheer et al., 2011; Kumar & Kanwar, 2012).

8.2 Submerged Fermentation This type of fermentation involves cultivation of microorganisms in a liquid medium. Most of the industrial lipase productions are done using this process. This involves growing selected microorganisms in a closed chamber containing a rich broth of nutrients and high amount of oxygen. As the microorganisms breakdown the nutrients, the enzyme is released in the liquid medium. Compared to SSF, SmF is used for many industrial applications

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and large scale productions. Fermentation process takes place in conical flask to large vessels of up to 1000 m3 (Selvam, 2011). There are various modes of fermentation with respect to the supply of nutrients for the growth of microorganisms: batch, fed-batch, repeated-batch, continuous modes. The choice of the fermentation modes depends on the characteristics of the product of interest (Bonugli-Santos et al., 2015). Fed-batch and continuous fermentation processes are more common. In fed-batch, the sterilized nutrients are added to the fermenter during the growth of the biomass. In continuous mode, sterilized liquid medium is fed into the fermenter at the same flow rate as the fermentation broth leaves the system (Selvam, 2011). Lipolytic enzyme from Thermos thermophilus HB27 is produced in a 5 L laboratory Table 5 Advantages and Disadvantages of Solid-State and Submerged Fermentation Solid-State Fermentation Submerged Fermentation Advantages

Disadvantages

Advantages

Higher volumetric productivity

Heat and mass transfer balances

Process takes place Yield is low compared to SSF in aseptic conditions

Low production costs and easy downstream processing

Scale up in industrial Mixing and level agitation is possible

Risk of contamination is high

Lowers catabolic repression

Easy availability Parameters of high of industrial enzyme yield and productivities need to equipments be carefully optimized

Not eco-friendly. High volumes of pollutants are produced

Extended stability Determination of of products biomass is difficult

Value-added conversion of biomass

–

Use of numerical – agricultural wastes as substrates

Ease of control of physical and chemical parameters

Disadvantages

High cost of technology and skilled person are required for the operation

Soluble substrates High energy are used consuming Ease of inoculation and continuous process

Limitation of soluble oxygen

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scale stirred tank and airlift bioreactor to determine the suitable reactor system for the production. Stirred tank bioreactor showed 2.3-fold higher productivity compared to airlift bioreactor. The study revealed that lipase production in stirred tank was less dependent on aeration than agitation based on relationship with mass transfer coefficient (Selvam, 2011). Production of lipase is influenced by the enzyme production and purification conditions (Table 5). The lipase produced through SSF and SmF was compared and the results differed in specific activity, heat stability, and fatty acid specificity (Bonugli-Santos et al., 2015).

9. CONCLUSION The recent research on efficient production of lipase is focused with marine microorganisms as a source because the commercial applications of lipase are increasing worldwide. The production of lipases from marine sources was proven to be effective in comparison to other terrestrial sources. This is because of their activity in extreme conditions. Production through SmF is preferred when it comes to large scale production due to many reasons like ease of availability of scale up equipment and periodic monitoring of physiochemical conditions for the growth of the microorganisms although SSF results in higher yield. Furthermore, isolation of novel species for production and purification of lipases which can retain its activity in broader range of pH, temperature, etc. is needed. With respect to SSF, more focus on scaling up of the process for large scale production and enhancing the technologies for monitoring the physiochemical parameters are required as the yield is higher. Bioprospection for such novel species and improving the techniques for better production and purification might help in easy availability, eco-friendly, and cost-effective solutions.

ACKNOWLEDGMENTS The authors sincerely thank Director Indian Institute of Technology Hyderabad for their continued encouragement and support. D.S.K. thanks Anup Ashok for helping in organizing chapter and SEED GRANT IITH-2014 for financial support.

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CHAPTER FIVE

Sequential Optimization Methods for Augmentation of Marine Enzymes Production in Solid-State Fermentation: L-Glutaminase Production a Case Study T. Sathish*,1,2, K.B. Uppuluri†, P. Veera Bramha Chari{, D. Kezia§ *Bioengineering and Environmental Centre, Indian Institute of Chemical Technology, Hyderabad, India † Bioprospecting Laboratory, School of Chemical and Biotechnology, SASTRA University, Thanjavur, India { Krishna University, Machilipatnam, India § Center for Biotechnology, Andhra University, Visakhapatnam, India 2 Corresponding author: e-mail addresses: [email protected]; [email protected]

Contents 1. Introduction 2. L-Glutaminase Production a Case Study 2.1 Experimentation 2.2 Observations 3. Conclusions References

96 97 97 102 113 113

Abstract There is an increased L-glutaminase market worldwide due to its relevant industrial applications. Salt tolerance L-glutaminases play a vital role in the increase of flavor of different types of foods like soya sauce and tofu. This chapter is presenting the economically viable L-glutaminases production in solid-state fermentation (SSF) by Aspergillus flavus MTCC 9972 as a case study. The enzyme production was improved following a three step optimization process. Initially mixture design (MD) (augmented simplex lattice design) was employed to optimize the solid substrate mixture. Such solid substrate mixture consisted of 59:41 of wheat bran and Bengal gram husk has given higher amounts of L-glutaminase. Glucose and L-glutamine were screened as a finest additional carbon and nitrogen sources for L-glutaminase production with help of Plackett– Burman Design (PBD). L-Glutamine also acting as a nitrogen source as well as inducer for secretion of L-glutaminase from A. flavus MTCC 9972. In the final step of optimization various environmental and nutritive parameters such as pH, temperature, moisture 1

Current address: Andaman and Nicobar Centre for Ocean Science and Technology, National Institute of Ocean Technology, Port Blair 744103, India.

Advances in Food and Nutrition Research, Volume 78 ISSN 1043-4526 http://dx.doi.org/10.1016/bs.afnr.2016.06.003

#

2016 Elsevier Inc. All rights reserved.

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content, inoculum concentration, glucose, and L-glutamine levels were optimized through the use of hybrid feed forward neural networks (FFNNs) and genetic algorithm (GA). Through sequential optimization methods MD–PBD–FFNN–GA, the L-glutaminase production in SSF could be improved by 2.7-fold (453–1690 U/g).

1. INTRODUCTION The environmental conditions of marine microorganisms are harsh and diverse; they can be hot or cold, acidic or basic, pressurized, saline, or mineral rich as found in deep ocean hydrothermal vents, polar oceans, and extremely saline water body. The extremophiles that survive in these diverse harsh conditions may produce enzymes that help in their survival (Chandrasekaran & Kumar, 2010). The marine enzymes are relatively stable and active than the corresponding enzymes from the terrestrial sources (Zhang & Kim, 2010). The major characteristics of marine microbial enzymes are salt tolerance, hyperthermostability, barophilicity, and cold adaptivity. L-Glutaminase (L-glutamine amidohydrolase, EC 3.5.1.2) is the enzyme deamidating L-glutamine. L-Glutaminase has many therapeutic and industrial applications. It is a potent anticancer agent and it acts as a flavor-enhancing agent in fermented foods by increasing the glutamic acid content and thereby imparting a palatable and enhanced taste (Kashyap, Sabu, Pandey, Szakacs, & Soccol, 2002; Sabu, Keerthi, Kumar, & Chandrasekaran, 2000). The enzyme can also be used as biosensor for monitoring glutamine levels in mammalian and hybridoma cell cultures (Katikala, Bobbarala, Tadimalla, & Guntuku, 2009). Salt tolerant and heat stable glutaminase from marine microbes are favored in food industry. Extracellular L-glutaminase is reported to be produced from marine yeasts (eg, Zygosaccharomyces rouxii) (Kashyap et al., 2002), marine fungi (eg, Beauveria sp.) (Sabu et al., 2000), and marine bacterium (eg, Pseudomonas sp.) (Kumar & Chandrasekaran, 2003). Production of marine microbial enzymes can be done using batch, continuous, or fed-batch fermentation. Batch process can be done by submerged fermentation (Estrada-Badillo & Ma´rquez-Rocha, 2003) and solid-state fermentation (SSF) (Kashyap et al., 2002). Production of L-glutaminase in SSF is an economically viable process (Sathish, Laxmi, Rao, Brahmaiah, & Prakasham, 2008). The selected solid substrate and cultural conditions play a vital role in the enzymes production in SSF. Optimization of various cultural conditions such as pH, temperature, moisture content, inoculum size and age of

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inoculum, and incubation time is vital for the production of microbial enzymes in SSF. Interactive effects of these factors have significant effect on the production of microbial products. The traditionally method of optimization is “one factor at a time” method, where only one parameter is optimized and all other parameters are kept constant. Though this method is simple, it lacks the interactive effects. To account the interactive effects of selected fermentation conditions, statistical methods such as factorial designs, orthogonal designs (Taguchi method), and response surface methodology are evolved (Sathish & Prakasham, 2010). Along with optimization methods, screening designs such as two-factorial designs and Plackett–Burman designs (PBDs) were also evolved, to select the best suitable component from a huge variety of choices with less number of experiments. Due to the availability of various software’s and literature on such statistical methods, many researchers are employing these designs in their work. Each method has some particular limitations as well as advantages. The assumption of mathematical models from linear to quadratic polynomials is the major limiting factor of these designs. To overcome these problems, artificial neural networks (ANNs), genetic algorithms (GAs), and particle swarm optimization are used to solve the nonlinear problems (Sathish & Prakasham, 2010). In this study, the production of L-glutaminase by Aspergillus flavus MTCC 9972 was improved by optimizing the various conditions by using statistical methods (MD, PBD) and artificial intelligence methods (ANN, GA). This case study represents a good example on how the application of combined statistical methods can help to optimize the enzyme production and obtain higher enzyme yields.

2. L-GLUTAMINASE PRODUCTION A CASE STUDY 2.1 Experimentation Marine fungus A. flavus MTCC 9972 isolated from coastal waters of Bay of Bengal at Visakhapatnam, A.P., India was used in this study. The spore solution prepared from 72 h culture was used as an inoculum. Different agroindustrial materials such as wheat bran (WB), rice husk (RH), rice bran (RB), corn cobs (CCs), sugar cane bagasse (SCB), red gram husk (RgH), green gram husk (GGH), Bengal gram husk (BeH), black gram husk (BgH), ground nut oil cake (GC), and coconut oil cake (CoC) were procured from the local market. The pineapple waste (PW) was obtained from the local fruit market and juice shops. The spent coffee (SC) and spent tea (ST) waste were collected from the canteen and home. In a different

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250 mL flask 10 g of processed substrate was taken individually, moisturized with 10 mL of distilled water and sterilized. The sterile substrates were initially received 1 mL of spore suspension (1
108 spores per mL) as inoculum. The content of flasks was thoroughly mixed and incubated at 28°C in a slanting position to get maximum surface area. Fermentation was carried up to 48 h. The crude enzyme was extracted by simple contact method as described by Sathish et al. (2008). L-Glutaminase activity was determined using L-glutamine as substrate (Imada, Igarasi, Nakahama, & Isono, 1973). Enzyme yield was expressed as units per gram of dry substrate (U/g). Mixture design (MD) was used to study the combined substrates impact on L-glutaminase production and optimized their concentrations to attain higher amounts of enzyme yield. Three best substrates suitable for fungal growth as well as enzyme production were chosen. In MD, the total proportions of the different factors must be 100% that is 10 g using three solid substrates. A 10 run augmented simplex lattice design was used in the present study. Table 1 shows the MD along with obtained L-glutaminase yield corresponding to the substrate mixture. Table 1 Mixture Design Along with Observed and Predicted L-Glutaminase Yield (U/g) L-Glutaminase Activity (U/g) Wheat Green Gram Bengal Gram S. No. Bran (g) Husk (g) Husk (g) Observed Predicted Error

1

1 (10)

0 (0)

0 (0)

453

454.57

1.57

2

0 (0)

1 (10)

0 (0)

388

385.66

2.34

3

0 (0)

0 (0)

1 (10)

402

401.94

0.06

4

0.5 (5)

0.5 (5)

0 (0)

436

435.24

0.76

5

0.5 (5)

0 (0)

0.5 (5)

502

503.52

1.52

6

0 (0)

0.5 (5)

0.5 (5)

406

403.61

2.39

7

0.6667 (6.7)

0.1667 (1.65)

0.1667 (1.65)

496

492.07

3.93

8

0.1667 (1.65)

0.6667 (6.7)

0.1667 (1.65)

428

435.79

7.79

9

0.1667 (1.65)

0.1667 (1.65)

0.6667 (6.7)

463

463.98

0.98

10

0.3333 (3.35)

0.3333 (3.35)

0.3333 (3.30)

494

491.57

2.43

The real values are presented in braces. Predicted values were calculated by special cubic model (Eq. 7).

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The analysis of MD varies from the response surfaces models because of the constraint Σxi ¼ 1. Linear to cubic models were used to determine the best blend of components for optimal medium. Y¼

p X

βi xi Linear model

(1)

i¼1

Y¼

p X

βi xi +

p XX

Y¼

p X

βi xi +

p XX

βij xi xj +

i