Marine Biotechnology I (Advances in Biochemical Engineering Biotechnology Vol 96)

96 Advances in Biochemical Engineering/Biotechnology Series Editor: T. Scheper

Editorial Board: W. Babel · I. Endo · S.-O. Enfors · A. Fiechter · M. Hoare · W.-S. Hu B. Mattiasson · J. Nielsen · H. Sahm · K. Schügerl · G. Stephanopoulos U. von Stockar · G. T. Tsao · C. Wandrey · J.-J. Zhong

Advances in Biochemical Engineering/Biotechnology Series Editor: T. Scheper Recently Published and Forthcoming Volumes

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With contributions by G. Antranikian · S. Barindra · G. Barnathan · J.-P. Bergé · C. Bertoldo G. Debashish · F. Guérard · M. Joydeep · M. Lalk · Y. Le Gal U. Lindequist · S. Malay · T. Matsunaga · H. Miyashita · T. Schweder D. Sellos · H. Takeyama · C. E. Vorgias · H. Yokouchi

123

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ISSN 0724-6145 ISBN-10 3-540-25659-8 Springer Berlin Heidelberg New York ISBN-13 978-3-540-25659-5 Springer Berlin Heidelberg New York DOI 10.1007/b135780

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Series Editor Prof. Dr. T. Scheper Institute of Technical Chemistry University of Hannover Callinstraße 3 30167 Hannover,Germany [email protected]

Volume Editors Prof. Dr. Roland Ulber

Prof. Yves Le Gal

Fachbereich Maschinenbau und Verfahrenstechnik Technische Universität Kaiserslautern Gottlieb-Daimler-Str., Gebäude 44/519 67663 Kaiserslautern, Germany [email protected]

Station de Biologie Marine Muséum National d’Histoire Naturelle Place de la Croix, BP 225 29182 Concarneau cedex, France [email protected], [email protected]

Editorial Board Prof. Dr. W. Babel

Prof. Dr. M. Hoare

Section of Environmental Microbiology Leipzig-Halle GmbH Permoserstraße 15 04318 Leipzig, Germany [email protected]

Department of Biochemical Engineering University College London Torrington Place London, WC1E 7JE, UK [email protected]

Prof. Dr. S.-O. Enfors

Prof. Dr. I. Endo

Department of Biochemistry and Biotechnology Royal Institute of Technology Teknikringen 34, 100 44 Stockholm, Sweden [email protected]

Faculty of Agriculture Dept. of Bioproductive Science Laboratory of Applied Microbiology Utsunomiya University Mine-cho 350, Utsunomiya-shi Tochigi 321-8505, Japan [email protected]

VI

Editorial Board

Prof. Dr. A. Fiechter

Prof. Dr. J.-J. Zhong

Institute of Biotechnology Eidgenössische Technische Hochschule ETH-Hönggerberg 8093 Zürich, Switzerland ae.fi[email protected]

State Key Laboratory of Bioreactor Engineering East China University of Science and Technology 130 Meilong Road Shanghai 200237, China [email protected]

Prof. W.-S. Hu Chemical Engineering and Materials Science University of Minnesota 421 Washington Avenue SE Minneapolis, MN 55455-0132, USA [email protected]

Prof. J. Nielsen Center for Process Biotechnology Technical University of Denmark Building 223 2800 Lyngby, Denmark [email protected]

Prof. Dr. B. Mattiasson Department of Biotechnology Chemical Center, Lund University P.O. Box 124, 221 00 Lund, Sweden [email protected]

Prof. Dr. H. Sahm Institute of Biotechnolgy Forschungszentrum Jülich GmbH 52425 Jülich, Germany [email protected]

Prof. Dr. G. Stephanopoulos Department of Chemical Engineering Massachusetts Institute of Technology Cambridge, MA 02139-4307, USA [email protected]

Prof. Dr. G. T. Tsao Director Lab. of Renewable Resources Eng. A. A. Potter Eng. Center Purdue University West Lafayette, IN 47907, USA [email protected]

Prof. Dr. K. Schügerl Institute of Technical Chemistry University of Hannover, Callinstraße 3 30167 Hannover, Germany [email protected]

Prof. Dr. U. von Stockar Laboratoire de Génie Chimique et Biologique (LGCB), Départment de Chimie Swiss Federal Institute of Technology Lausanne 1015 Lausanne, Switzerland urs.vonstockar@epfl.ch

Prof. Dr. C. Wandrey Institute of Biotechnology Forschungszentrum Jülich GmbH 52425 Jülich, Germany [email protected]

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Preface

Oceans, which occupy up to two thirds of the surface of our planet, were not really approached from scientific point of view until the second half of the 19th century and even the 20th with regard to microbial and unicellular life. Today, the importance of marine biodiversity has been fully recognized. It is, indeed, one of the aspects which, over the two past decades, have made a major contribution to our knowledge and vision of the living planet. Marine organisms make up a very large collection of living beings, of which the number of species is likely to exceed that of land species by several orders of magnitude. However, the major interest in marine living systems is mainly linked to the exceptional diversity of molecular structures, of chemical interactions reflecting a long evolutionary history and also the fact that marine life spreads within multiple and complex three-dimensional spaces. From a very practical and trivial point of view, marine life can be seen as a huge reservoir of molecules which enable marine organisms to defend themselves, to communicate and, more generally, to survive and thrive. In a very broad sense, marine biotechnologies can be understood as being the various techniques of managing marine living systems for the profit of mankind. The domain covered by marine biotechnologies is vast and ranges over various overlapping disciplines, from developmental biology to the chemistry of natural substances and bioprocess engineering. Not all these fields, however, are ready for practical and industrial applications. Biomass from fishing or the aquaculture industry is, in fact, complex, geographically and seasonally dependent mixtures of compounds requiring adapted purification procedures. Furthermore, many “natural substances”, for which we do not know any terrestrial counterparts, and are, therefore, of the greatest interest, only exist in tiny amounts in rare biological species and their exploitation is likely to call for costly synthetic procedures. Originally, marine life was used essentially as a provider of biomass directly and indirectly for food. However, today, in the global context of the standstill of world fisheries, common sense requires us to upgrade and exploit the actual biomass better. This clearly means developing techniques for identifying the source of raw materials suitable for specific biomolecules, the design of processes and their scaling up from pilot plant to production processes.

X

Preface

Marine natural products not only display novel characteristics but also complexity in terms of chemical structures. Isolation and structure elucidation represents only the tip of the iceberg. The question of the functionality of new isolated molecules within the perspective of challenging major public health and environmental problems is crucial. In this domain, ecological and evolutionary approaches should help the classical screening systems for determining the right target systems. In addition, a better understanding of the complex interactions between macro and micro organisms is necessary for us to be able to use these resources for industrial purposes. Thus, more and more groups are focussing on the field of bioprocess engineering and downstream processing in marine biotechnology. This volume of Advances in Biochemical Engineering/Biotechnology illustrates several topics in line with the following broad objectives: thinking of marine biotechnology as the controlled production and use of marine organisms and molecules for useful purposes, firstly by exploring aspects of marine biodiversity and exploitation of biomass, then considering the identification, production and processing of marine products. Kaiserslautern and Concarneau, August 2005

Roland Ulber and Yves Le Gal

Contents

Screening for New Metabolites from Marine Microorganisms T. Schweder · U. Lindequist · M. Lalk . . . . . . . . . . . . . . . . . . .

1

Fatty Acids from Lipids of Marine Organisms: Molecular Biodiversity, Roles as Biomarkers, Biologically Active Compounds, and Economical Aspects J.-P. Bergé · G. Barnathan . . . . . . . . . . . . . . . . . . . . . . . . . .

49

Fish and Shellfish Upgrading, Traceability F. Guérard · D. Sellos · Y. Le Gal . . . . . . . . . . . . . . . . . . . . . . 127 Marine Microalgae T. Matsunaga · H. Takeyama · H. Miyashita · H. Yokouchi . . . . . . . . 165 Marine Enzymes G. Debashish · S. Malay · S. Barindra · M. Joydeep . . . . . . . . . . . . 189 Extreme Environments as a Resource for Microorganisms and Novel Biocatalysts G. Antranikian · C. E. Vorgias · C. Bertoldo . . . . . . . . . . . . . . . . 219 Author Index Volumes 51–97 . . . . . . . . . . . . . . . . . . . . . . . 263 Subject Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 285

Contents of Volume 97 Marine Biotechnology II Volume Editors: Yves Le Gal, Roland Ulber ISBN: 3-540-25669-5

Aquaculture of “Non-Food Organisms” for Natural Substance Production G. Liebezeit Bioprocess Engineering Data on the Cultivation of Marine Prokaryotes and Fungi S. Lang · M. Hüners · V. Lurtz Downstream Processing in Marine Biotechnology K. Muffler · R. Ulber Marine Pharmacology: Potentialities in the Treatment of Infectious Diseases, Osteoporosis and Alzheimer’s Disease M.-L. Bourguet-Kondracki · J.-M. Kornprobst Asymmetric Total Synthesis of Complex Marine Natural Products J. Hassfeld · M. Kalesse · T. Stellfeld · M. Christmann Seafood Allergy: Lessons from Clinical Symptoms, Immunological Mechanisms and Molecular Biology K. H. Chu · C. Y. Tang · A. Wu · P. S. C. Leung

Adv Biochem Engin/Biotechnol (2005) 96: 1–48 DOI 10.1007/b135781 © Springer-Verlag Berlin Heidelberg 2005 Published online: 24 August 2005

Screening for New Metabolites from Marine Microorganisms Thomas Schweder (u) · Ulrike Lindequist · Michael Lalk Institut für Marine Biotechnologie, W.-Rathenau-Str. 49, 17489 Greifswald, Germany [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2

2 2.1 2.2 2.3 2.4 2.4.1 2.4.2 2.5

Sequencing of the Genomes of Marine Microorganisms . . . . . . . . . . . Completed and Ongoing Marine Sequencing Projects . . . . . . . . . . . . Analysis of Marine Microbial Diversity . . . . . . . . . . . . . . . . . . . . Environmental Genomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . Functional Genome Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . Proteome Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Transcriptome Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genome Sequencing and Identification of New Antimicrobial Compounds

3 3 18 18 22 22 23 24

3 3.1 3.2 3.3 3.4 3.5 3.6 3.7

Screening for New Metabolites . . . . . . . . . . . . . . . . . Alternative Cultivation Methods . . . . . . . . . . . . . . . . Preparation of Materials for Screening . . . . . . . . . . . . . Chemical and Physicochemical Screening . . . . . . . . . . . Biological Screening . . . . . . . . . . . . . . . . . . . . . . . High-Throughput Screening, Automation, Data Management Metabolome Analysis Techniques . . . . . . . . . . . . . . . Examples for Metabolites from Marine Microorganisms . . .

26 26 27 28 28 30 31 32

4

Application of Proteomics for Target Analyses of Antibacterial Compounds

41

5

Influence of Cultivation Conditions on Metabolite Production . . . . . . .

41

6

Outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

42

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

42

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. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

. . . . . . . .

Abstract This article gives an overview of current analysis techniques for the screening and the activity analysis of metabolites from marine (micro)organisms. The sequencing of marine genomes and the techniques of functional genomics (including transcriptome, proteome, and metabolome analyses) open up new possibilities for the screening of new metabolites of biotechnological interest. Although the sequencing of microbial marine genomes has been somewhat limited to date, selected genome sequences of marine bacteria and algae have already been published. This report summarizes the application of the techniques of functional genomics, such as transcriptome analysis in combination with high-resolution two-dimensional polyacrylamide gelelectrophoresis and mass spectrometry, for the screening for bioactive compounds of marine microorganisms. Furthermore, the target analysis of antimicrobial compounds by proteome or transcriptome analysis of bacterial model systems is described. Recent high-throughput screening

2

T. Schweder et al.

techniques are explained. Finally, new approaches for the screening of metabolites from marine microorganisms are discussed. Keywords Functional genomics · Proteome · Metabolome · Natural compounds · High-throughput screening

1 Introduction The tremendous biochemical diversity of marine microorganisms and their biotechnological potential is becoming more and more recognized, not only by microbiologists but also by the pharmaceutical industry. Several new companies focus on the discovery of more effective drugs based on natural products of marine microorganisms. In recent years, the improvement of screening technologies has yielded a considerable number of potential new drug candidates and other metabolites from microorganisms of marine ecosystems. The physiological investigation of marine natural products has shown that many of these compounds function as signal transducers and thus regulate complex processes within marine living societies. It is supposed that these structures play an important role in the inhibition of physiological processes of potential competitors. This offers a promising potential for the exploration of new drugs against critical pathogenic microorganisms. Most of the marine compounds that have been successfully screened and structurally elucidated so far originate from microorganisms, especially bacteria. Several studies have demonstrated that natural products isolated from higher marine organisms like marine invertebrates are very frequently of bacterial origin. However, these bacteria, which are for example in a symbiotic association with higher organisms, usually cannot be cultivated alone in a pure culture. Their growth depends directly on the activity of their hosts. Furthermore, only a minority of free-living marine microorganisms has been identified and can be cultivated so far. The knowledge on such marine microorganisms is very limited compared to those microorganisms that can be easily cultured under laboratory conditions. The taxonomical identification of marine microorganisms in general is still in its infancy. The focus on the physiology and the potential of bioactive substances of non-cultivable marine microorganisms is an important challenge at present and for the future. The estimated taxonomical diversity of marine microorganisms in general indicates the powerful potential of novel bioactive substances produced in aquatic ecosystems. It has been shown that marine bacteria, which are the predominant microorganisms in the marine ecosystems, produce bioactive substances that are different from known compounds from terrestrial bacte-

Screening for New Metabolites from Marine Microorganisms

3

ria. Also, metabolites from facultative and obligate marine fungi often have structures unlike those of their terrestrial counterparts [1]. In the past, a successful cultivation was usually the prerequisite for the screening and final application of a new natural compound. However, during the last few years promising techniques have been developed that allow the screening and presumably also application of biological activities, not only of cultivable marine microorganisms but also of those organisms that cannot be cultivated at present. These techniques are the focus of this article.

2 Sequencing of the Genomes of Marine Microorganisms The genome sequencing of an organism gives the blueprint of its life. This blueprint establishes the basis for a comprehensive view of the cellular physiology. Knowing the sequence of all genes does not only allow the identification of protein functions but also makes it possible to explore the complexity of the cellular organization of an organism. Elucidation of the structural organization of sequenced genomes has led to new insights into the physiological capacity of these organisms. This opens up new possibilities for the exploration of genes that are involved in pathways responsible for the synthesis of metabolites of biotechnological interest. However, the functions of the majority of genes are still unknown. Understanding these functions will be a major challenge for the next decades. In this field we are only at the beginning. 2.1 Completed and Ongoing Marine Sequencing Projects A prerequisite for the sequencing of the whole genome of an organism is usually the cultivation of the cells in a pure culture. It is estimated that about 1–10% of the microbial diversity on earth has been identified [2]. However, the majority of the taxonomically classified microorganisms are of terrestrial origin. The considerably smaller number of identified marine microbial species is one of the reasons why most of the microbial genomes that have been sequenced so far originate from terrestrial microorganisms. Only few marine genome projects have been started and finished up to now. The first completely sequenced genomes of marine bacteria were that of the methanogenic bacterium Methanococcus jannaschii [3] and that of the cyanobacterium Synechocystis spp. [4]. A selection of completed and ongoing genome sequencing projects dealing with marine microorganisms is given in Table 1.

Aerobic hyperthermophilic archaeon 1.669 growing at temperatures up to 100 ◦ C Oil-degrading (hydrocarbonoclastic), 3.12 surfactant-producing marine bacterium

Aeropyrum pernix

Alcanivorax borkumensis

Photosynthetic, anaerobic, green-sulfur bacterium, thermophile

Chlorobium tepidum TLS

TIGR, USA

TIGR, Center of Marine Biotechnology, USA

2.1

Thermophilic anaerobic bacterium from hydrothermal environments

Carboxydothermus hydrogenoformans

Competence Network Bielefeld, Germany

7.212 Kazusa DNA Research (1 chromosome of 6.414 Mb and Institute, Japan six plasmids of 408.101 bp, 186.614 bp, 101.965 bp, 55.414 bp, 40.340 bp and 5.584 bp)

2.155

[5]

Reference

[10]

[9]

[8]

[7]

National Institute of [6] Technology and Evaluation (NITE), Japan

Diversa, San Diego/USA

Institution

Anabaena sp. strain PCC 7120 Cyanobacterium

1.551

Hyper-thermophilic marine bacterium (85 – 95 ◦ C)

Aquifex aeolicus VF5

Genome size (Mb)

Properties

Organism

Complete genome sequences of marine microorganisms

Table 1 Selection of completed and ongoing marine genome sequencing projects

4 T. Schweder et al.

Properties Arctic bacterium Antarctic archaechon (one of two cold-adapted archaeans to be sequenced) Methanogenic archeon Extremophile, grows near hydrothermal vents at pressures of more than 200 atm and optimum temperature of 85 ◦ C Hyperthermophile methaneproducing archaeon at the base of black smokers (temperatures near 100 ◦ C) Marine archaeon, living as an obligate symbiont on another microbe (Ignicoccus) in undersea vents and hot springs (one of the smallest genomes of any sequenced microbe) Bacillus-related deep sea sediment bacterium, tolerates extremely saline and alkaline environments

Organism

Colwellia psychroerythraea 34H

Methanococcoides burtonii

Methanococcus jannaschii DSM 2661

Methanopyrus kandleri AV19

Nanoarchaeum equitans

Oceanobacillus iheyensis HTE831

Table 1 (continued)

3.6

0.491

1.695

1.66

2.8–3

5.3

Size (Mb)

Japan Marine Science and Technology Center

Diversa Corporation, San Diego/USA

Fidelity Systems, Inc, USA

TIGR, USA

Joint Genome Institute, USA

TIGR, USA

Institution

[16]

[15]

[14]

[13]

[12]

[11]

Reference

Screening for New Metabolites from Marine Microorganisms 5

Properties Marine cyanobacterium, one of the most abundant photosynthetic organisms on earth Low-light-adapted Low-light-adapted ecotype (MIT 9313) in deeper waters with less sunlight Highly light-adapted ecotype (MED4), near the ocean surface Facultatively aerobic nitratereducing hyperthermophilic crenarchaeon, T(opt) = 100 ◦ C, isolated from a boiling marine water hole Archaeon, growing close to hot springs 3500 m deep in the southeast Pacific (optimally at 103 ◦ C and 200 atmospheres) Archaeon found in marine sand surrounding sulfurous volcanoes, optimal growth at temperatures above 100 ◦ C, highly resistant to radiation

Organism

Prochlorococcus marinus SS120

Prochlorococcus marinus MIT9313

Prochlorococcus marinus subsp. pastoris CCMP1378 (MED4)

Pyrobaculum aerophilum DC3000

Pyrococcus abyssi GE5

Pyrococcus furiosus DSM 3638

Table 1 (continued)

1.908

1.765

2.2

1.658

2.411

1.86

Size (Mb)

University of Utah & University of Maryland, USA

Genoscope, Frankreich

University of California & California Institute of Technology, USA

Joint Genome Institute, USA

Joint Genome Institute, USA

Genoscope and Station Biologique de Roscoff, France

Institution

[21]

[20]

[19]

[18]

[18]

[17]

Reference

6 T. Schweder et al.

Marine planctomycete from the Baltic Sea, emits a reddish sunscreen to protect itself from sunlight Aerobic thermoacidophilic crenarchaeon, optimal growth at 80 ◦ C (hot springs), at low pH Marine unicellular cyanobacterium, photosynthetic, motile, in the open ocean Unicellular aquatic, photosynthetic cyanobacterium Thermophilic unicellular cyanobacterium (hot springs)

Rhodopirellula baltica (formerly Pirellula sp. strain 1)

Sulfolobus tokodaii 7

Synechococcus WH8102

Synechocystis PCC 6803

Thermosynechococcus elongatus BP-1

Archaeon, optimal growth at 106 ◦ C (in hydrothermal vents at the bottom of the Atlantic Ocean)

Hyperthermophilic archaebacterium near hydrothermal vents, extremophile (optimum growth temperature 98 ◦ C)

Pyrococcus horikoshii OT3

Pyrolobus fumarii

Properties

Organism

Table 1 (continued)

2.594

3.573

2.4

2.695

7.145

1.8

1.739

Kazusa DNA Research Institute, Japan

Kazusa DNA Research Institute, Japan

Joint Genome Institute and Scripps Institute, USA

National Institute of Technology and Evaluation (NITE), Japan

Max Planck Institute of Marine Microbiology, Germany

Diversa, Celera Genomics, USA

National Institute of Technology and Evaluation, Japan

Size (Mb) Institution

[28]

[27]

[26]

[25]

[24]

[23]

[22]

Reference

Screening for New Metabolites from Marine Microorganisms 7

Marine bioluminescent bacterium, specific symbiont in the lightemitting organs of certain species of squids and fishes Halophilic seawater pathogen, causes wound infections, gastroenteritis

Vibrio fischeri ES114

Vibrio vulnificus

Integrated Genomics Inc, Univ. of Hawaii, USA

The Institute for Genomic Research (TIGR), Maryland/USA

Institution

Salmonid bacterial pathogen

Aeromonas salmonicida subsp. salmonicida

Bacteriovorax marinus SJ Marine predatory bacterium (former salt water Bdellovibrio sp.) that parasitises a wide range of Gram negative bacteria

Enterotoxic aeromonad, fish bacterial pathogen

Aeromonas hydrophila

3.431

4.7

No data

[32]

[31]

[30]

[29]

Reference

Sanger Institute, University of Maryland, Morgan State University, USA

[32]

National Research Council, [32] Institute for Marine Biosciences, Canada

TIGR, University of Maryland, USA

Genomes and chromosomes of marine microorganisms in progress

5.261 (2 chromosomes of Yang-Ming University, 3.355 Mb and 1.857 Mb, one Taiwan plasmid of 48.508 bp)

4.136

Extremely thermophilic eubacterium 1.861 growing up to 90 ◦ C

Thermotoga maritima MSB 8

Size (Mb)

Properties

Organism

Table 1 (continued)

8 T. Schweder et al.

Properties Green sulfur bacterium Green sulfur bacterium Green sulfur bacterium Phototrophic gliding filamentous bacterium of hot springs Psychrophilic crenarchaeon that inhabits a marine sponge Diazotropic marine cyanobacterium, isolated from the tropical Atlantic and Pacific Oceans Extremely radiation-resistant and slightly thermophilic bacterium from hot spring

Organism

Chlorobium limicola DSMZ 245(T)

Chlorobium phaeobacteroides DSMZ 266(T) and MN1

Chlorobium vibrioforme f. thiosulfatophilum DSMZ 265(T)

Chloroflexus aurantiacus J-10-fl

Cenarchaeum symbiosum

Crocosphaera watsonii WH8501

Deinococcus geothermalis DSM11300

Table 1 (continued)

No data

4.0

3.7

3.0

2.4

2.4

2.4

Joint Genome Institute, USA

Joint Genome Institute, Woods Hole Oceanographic Institution, USA

Monterey Bay Aquarium Research Institute, USA

Joint Genome Institute, USA

Joint Genome Institute, USA

Joint Genome Institute, USA

Joint Genome Institute, USA

Size (Mb) Institution

[32]

[32]

[32]

[32]

[32]

[32]

[32]

Reference

Screening for New Metabolites from Marine Microorganisms 9

Properties Originally isolated from marine mud of the Mediterranean Sea

Psychrophilic sulfate-reducing bacterium isolated from permanently cold Arctic marine sediments Anaerobic, sulfur-reducing, acetate-oxidizing bacterium from sea or fresh water Primary bacterial pathogen of channel catfish

Lives in the intestines of several kinds of surgeonfish off the Australian shore, up to 500 µm in size

Organism

Desulfobacterium autotrophicum HRM2

Desulfotalea psychrophila LSv54

Desulfuromonas acetoxidans

Edwardsiella ictaluri 93-146

Epulopiscium sp.

Table 1 (continued)

No data

No data

4.1

3.66

> 5.5

Size (Mb)

TIGR

Oklahoma University Health Science Center, Mississippi State University, USA

Joint Genome Institute, USA

Epidauros Biotechnologie AG, REGX, Germany

Göttingen Genomics Laboratory, Real Environmental Genomics (REGX, Max Planck Institute for Marine Microbiology, Bremen), Germany

Institution

[32]

[32]

[32]

[32]

[32]

Reference

10 T. Schweder et al.

Properties Fish pathogen (coldwater disease) in aquaculture

Extracellular polysaccharideproducing marine bacterium Extremely halophilic archaeon from the Dead Sea

Hyperthermophilic peptidefermenting sulfur archaebacterium from the sea floor of solfataric habitats Primary colonizers of surfaces in marine environments and in areas adjacent to hydrothermal vents Isolated from microaerobic zones of freshwater sediments

Organism

Flavobacterium psychrophilum

Hahella chejuensis 96CJ10356

Haloarcula marismortui ATCC43049

Hyperthermus butylicus

Hyphomonas neptunium ATCC 15444

Magnetospirillum gryphiswaldense

Table 1 (continued)

No data

2.7

1.9

2.7

7.0

No data

Size (Mb)

Max-Planck-Institute of Marine Microbiology, Max-Planck-Institute of Genetics, Germany

University of Georgia, TIGR, USA

Epidauros Biotechnologie AG, Germany, University of Copenhagen, Denmark

University of Maryland Biotechnology Institute, Institute for Systems Biology, USA

Korea Research Institute of Bioscience and Biotechnology

National Center for Cool and Cold Water Aquaculture, Integrated Genomics, USA

Institution

[33]

[32]

[32]

[32]

[32]

[32]

Reference

Screening for New Metabolites from Marine Microorganisms 11

Properties From microaerobic zones of freshwater sediments Thermophilic nitrogen fixing archaeon from submarine hydrothermal vents Archaeon from submarine hydrothermal vents Psychrophilic, H2 -using methanogenic archaeon from Ace Lake, Antarctica Acetotrophic methane-producing bacterium isolated from marine sediments Marine methylotroph

Aerobic marine bacterium that degrades and recycles complex carbohydrates

Organism

Magnetospirillum magnetotacticum MS-1, ATCC 31632

Methanococcus thermolithotrophicus

Methanococcus voltae

Methanogenium frigidum

Methanosarcina acetivorans

Methylophaga thalassica S1

Microbulbifer degradans 2-40

Table 1 (continued)

6.0

No data (1995-bp Plasmid)

4.1

2–2.5

No data

No data

4.5

Joint Genome Institute, USA

Integrated Genomics Inc, USA

Göttingen Genomics Laboratory, Germany

Univ. of New S. Wales, Australian Genome Research Facility, Australia Molecular Dynamics, UK

Molecular Dynamics, Integrated Genomics Inc, USA

Molecular Dynamics, UK Integrated Genomics, USA

Joined Genome Institute, USA

Size (Mb) Institution

[32]

[32]

[32]

[12]

[32]

[32]

[32]

Reference

12 T. Schweder et al.

Properties Bloom-forming toxic cyanobacterium Close relative of M. tuberculosis, fish and human pathogen

Anaerobic, thermophilic sulfurreducing bacterium, isolated from tubes of the deep-sea hydrothermal vent polychaete Alvinella pompejana Alpha purple bacterium, dominant microorganism in the ocean surface (bacterioplankton) Thermophilic hydrogen-oxidizing microaerophile from deep-sea hydrothermal vents

Organism

Microcystis aeruginosa PCC 7806

Mycobacterium marinum M

Nautilia sp. Am-H

Pelagibacter ubique HTCC1062

Persephonella marina

Table 1 (continued)

1.6

1.54

No data

6.739

4.8

Portland State Univ., TIGR, USA

Oregon State Univ., Diversa, Center of Marine Biotechnology, USA

Univ. of Delaware TIGR, USA

Sanger Institute, UK, University of Washington, USA, Institute Pasteur, France, Monash Univ., Australia, Univ. of Tennessee, USA

Institute Pasteur, France

Size (Mb) Institution

[32]

[32]

[32]

[34]

[32]

Reference

Screening for New Metabolites from Marine Microorganisms 13

Properties Antarctic psychrotolerant seawater bacterium

Actinobacterium, causative agent of bacterial kidney disease in salmonid fishes Isolated from eggs and accessory nidamental glands of squids Gram negative gamma proteobacterium from the Baltic Sea Antarctic Gram negative gamma proteobacterium Deep-sea barophilic Shewanella strain

Organism

Pseudoalteromonas haloplanktis

Renibacterium salmoninarum

Roseobacter sp. TM1040

Shewanella baltica OS1155

Shewanella frigidimarina NCMB400

Shewanella violacea DSS12

Table 1 (continued)

4.9

No data

No data

No data

No data

Two chromosomes of approx. 2700 and 800 kb

Size (Mb)

Kinki Univ., JAMSTEC, Keio Univ., Japan

Joint Genome Institute, USA

Joint Genome Institute, USA

Joint Genome Institute, USA

NWFSC, NCCCWA, Univ. of Washington, Integrated Genomics, USA

Genoscope, France

Institution

[32]

[32]

[32]

[32]

[32]

[35]

Reference

14 T. Schweder et al.

Properties Dimethylsulfoniopropionatedemethylating bacteria from marine environments Chlorophenol-degrading alpha Proteobacterium from seawater fjords in Alaska Blue-green microalgae from lakes with high salt concentrations, high nutrient-density Extremely thermophilic heterotrophic archaeon, occurring in heated sea flows Extremely thermophilic eubacterium near deep sea vents Autotrophic gamma proteobacterium from hydrothermal vent sites

Organism

Silicibacter pomeroyi DSS-3

Sphingopyxis alaskensis RB2256

Spirulina platensis

Thermococcus kodakaraensis KOD1

Thermotoga neapolitana ATCC49045

Thiomicrospira crunogena

Table 1 (continued)

No data

1.8

No data

No data

No data

4.4

Size (Mb)

Joint Genome Institute, USA

TIGR, USA

Kyoto Univ. Kwansei Gakuin Univ., Japan

Human Genome Center, Beijing, China

Joint Genome Institute, USA

Univ. of Georgia, TIGR, USA

Institution

[32]

[32]

[32]

[32]

[32]

[32]

Reference

Screening for New Metabolites from Marine Microorganisms 15

No data

Fish pathogen

Vibrio salmonicida

6.5

3.6–4.0

Marine filamentous cyanobacterium in tropical and subtropical ocean

Trichodesmium erythraeum IMS101

Size (Mb)

Uncultivated_Riftia pachyplila_endosymbiont Symbiont of deep sea tube worm near hydrothermal vents and hot springs

Properties

Organism

Table 1 (continued)

Univ. of Tromso, NORSTRUCT, Norway

Molecular Dynamics, Scripps Institution of Oceanography, Quorex, USA

Joint Genome Institute, Woods Hole Oceanographic Institute, USA

Institution

[32]

[32]

[32]

Reference

16 T. Schweder et al.

Screening for New Metabolites from Marine Microorganisms

17

Recently, the genomes of three different members of the cyanobacterium species Prochlorococcus marinus have been sequenced [36, 37]. These members of the genus Prochlorococcus dominate phytoplankton communities in most tropical and temperate open ocean ecosystems [38]. P. marinus is found in two main ecological forms: high-light-adapted genotypes in the upper part of the water column and low-light-adapted genotypes at the bottom of the illuminated layer [36]. Analysis of the genome sequences of the different P. marinus strains led to the identification of genes that play a role in determining the relative fitness of the ecotypes in response to key environmental variables [37]. Furthermore, the genome comparison allowed determination of a putative minimal gene set of this photoautotrophic bacterium [36]. Glöckner et al. (2003) reported the complete genome sequence of Pirellula sp. strain 1 (“Rhodopirellula baltica”), a marine representative of the globally distributed and environmentally important bacterial order Planctomycetales [39]. With 7.145 megabases, Pirellula sp. strain 1 has the largest circular bacterial genome sequenced so far. The genome sequencing indicated the presence of all genes required for heterolactic acid fermentation, key genes for the interconversion of C1 compounds, and 110 sulfatases, which were unexpected for this aerobic heterotrophic isolate. The first genome of an uncultivable marine bacterium, the symbiont of the deep sea tube worm Riftia pachyptila [40] is currently being sequenced by a Californian consortium (Horst Felbeck, personal communication). This uncultivable endosymbiotic bacterium seems to form a monoculture in the trophosome of R. pachyptila. The knowledge of the bacterial genome sequence together with molecular techniques such as the proteome analysis might help to find the explanation for this exclusive colonization by the microbial endosymbiont inside the trophosome of the tube worm. Furthermore, the functional genome analysis of the bacterial symbiont will lead to the identification of mechanisms and possibly natural products involved in establishing and maintaining the symbiosis with R. pachyptila. Another promising field is the genome analysis of marine phages [41]. Phages have evolved unique strategies that sustainably influence the cellular processes of bacteria. The directed isolation of marine phages and the determination of their genome sequence could help to establish new strategies for the inhibition of growth of pathogenic marine bacteria, but also of human pathogens. Such an approach has been undertaken by Liu et al. (2004) for Staphylococcus aureus phages [42]. The sequencing of 26 phage genomes led to the identification of 31 novel polypeptide families, the expression of which inhibited the growth of S. aureus. Furthermore, the analysis of potential targets of these polypeptides led to the determination of proteins essential for bacterial DNA replication. A subsequent screening for small molecule inhibitors of these targets allowed the isolation of chemical compounds that imitate the growth-inhibiting effect of selected phage polypeptides. Thus, this

18

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phage genomic approach could be a further strategy for the identification of new antibiotics to successfully combat infectious diseases. 2.2 Analysis of Marine Microbial Diversity The majority of marine microbial organisms cannot be cultured under artificial laboratory conditions and is thus not accessible for detailed taxonomical and physiological characterizations. However, advanced molecular techniques have altered the perspective on naturally occurring diversity and distribution of such marine microorganisms. Direct isolation of DNA from the environment makes it possible to identify bacterial species in different natural marine habitats without the cultivation of the microbial cells. This analysis is mainly based on the selective amplification of 16S rRNA gene sequences by application of primers of conserved regions of the 16S rDNA in combination with the polymerase chain reaction (PCR) [43]. Due to its highly conserved and variable sequence regions the 16S rDNA sequence is used as a phylogenetic marker. The 16S rDNA sequence allows the identification and taxonomical affiliation of different microbial strains from an environmental sample. This analysis is thus an important feature, which allows the identification of so far unknown organisms and gives valuable information on the biodiversity in marine habitats [44]. However, this technique gives only limited information on the distribution and physiology of the microorganisms in their natural environment. The fluorescence in situ hybridization (FISH) allows investigations on the structure of a population without PCR amplification of specific sequences or cultivation of the microorganisms [45]. FISH with rRNA specific probes is a technique that allows phylogenetic identification of bacteria in mixed assemblages. For these analyses epifluorescence or confocal laser scanning microscopy are applied [47, 48]. The sensitivity of the FISH 16S rRNA analysis technique is based on the high number of ribosomes per cell and that each ribosome contains one copy of the 16S rRNA. This is a kind of natural signal amplification system. 2.3 Environmental Genomics The FISH technique leads to a better understanding of the identity of selected microorganisms and their distribution and abundance in the appropriate marine habitats. However, information concerning the biological properties of these marine bacteria cannot be satisfyingly illuminated this way. In this respect a new strategy has been developed, which is called environmental genomics. Similar to the sequencing of genomes from cultivable microorganisms, chromosomal DNA is used to generate genomic libraries. By the

Screening for New Metabolites from Marine Microorganisms

19

environmental genomic strategy not only one genome is considered but all genomic sequences from one environmental sample. Large genomic DNA fragments are directly isolated from the environment and cloned into suitable host vector systems. Establishment of comprehensive gene libraries attempts to cover all genome sequences from an environmental sample, to gather as much information as possible on the biosynthetic machinery of a microflora [49]. The comprehensive coverage of the genomes from an environmental sample is also called metagenome analysis. This technique allows a more realistic understanding of prokaryotic biodiversity in a distinct marine habitat [50]. A prerequisite for establishing comprehensive genomic libraries from environmental samples is the availability of host vector systems allowing the stable propagation of large DNA fragments. For the cloning of large genome fragments usually fosmid [51] or bacterial artificial chromosome (BAC) [52, 53] vectors are used (Fig. 1). Fosmid or BAC vectors facilitate the conservation of large genomic fragments and thus open up the possibility to characterize not only the gene content but also the physiological potential of uncultivated microorganisms. The BAC system is based on the Escherichia coli single copy F factor plasmid. The E. coli F factor replicon allows for a copy number control of the clones so that they are maintained at one to two copies per cell. BAC vectors can carry DNA inserts of variable size with a maximum of 300 kb. Similar to the BAC system, the fosmid vectors also carry the E. coli

Fig. 1 Cloning strategies for the screening of enzymatic or metabolic activities

20

T. Schweder et al.

F factor replicon, which provides for their stable single copy number. Fosmid vectors allow a size selection of the cloned DNA fragments from 32 to 43 kb by packaging the DNA in λ-phage heads. Shizuya et al. (1992) developed the bacterial cloning system BAC for mapping and analysis of complex genomes [54]. Because of its high cloning efficiency and the stable maintenance of inserted DNA, the BAC system is able to facilitate the construction of DNA libraries of complex environmental genomic samples but also provides a comprehensive representation of the genome sequence of one organism. The stabilizing effect of BAC and also fosmid vectors is an important feature for the generation of comprehensive genome libraries since distinct regions of genomic DNA (e.g., coding for potential toxic proteins) can cause vector instabilities in high copy numbers. The ability to clone long stretches of DNA has become an important tool for genome analyses of uncultivated marine microorganisms (Fig. 1). Stein et al. [51] isolated large genomic fragments from a widely distributed and relatively abundant, but as yet uncultivated, group of prokaryotes, the planktonic marine archaea from marine picoplankton. By construction and analysis of a fosmid DNA library a 38.5-kbp recombinant clone could be identified, which contained an archaeal small subunit ribosomal DNA gene. A similar approach was used to identify genomic DNA fragments of the symbiotic marine archaeon Cenarchaeum symbiosum from the total DNA of the marine sponge Axinella mexicana [55]. An environmental genomic study of Beja et al. [56] led to the discovery of proteorhodopsin, a retinal-containing integral membrane protein that functions as a light-driven proton pump, in the genome of an uncultivated marine bacterium. This study indicated that photoactive proteorhodopsin is present in oceanic surface waters. The data of Beja et al. (2001) suggested that proteorhodopsin-based phototrophy is a globally significant oceanic microbial process. In another study Beja et al. [57] analyzed large genome fragments from microorganisms sampled from an Antarctic picoplankton population and compared them to those from deeper waters of the temperate North Pacific. This study was initiated to better characterize uncultivated planktonic crenarchaeotes, which are present in high abundance in Antarctic winter surface waters and deep ocean waters. For this purpose environmental genomic fragments were cloned into fosmid vectors and the inserts were sequenced. Analysis of the DNA insert of one Antarctic marine archaeon revealed differences in genome structure and content between archaea from Antarctic surface water and temperate deepwater. Analysis of the proteins encoded by the archaeon sequence from surface water and those derived from a deepwater planktonic crenarchaeote revealed many typical archaeal proteins but also several proteins that have not been detected in archaea so far. Furthermore, a comparison of closely related archaea originating from a single population revealed significant genomic sequence differences that were not evident from

Screening for New Metabolites from Marine Microorganisms

21

the 16S rRNA sequence analysis. Beja et al. [57] concluded that considerable functional diversity may exist within single populations of coexisting microbial strains, even those with identical 16S rRNA sequences. Metabolic features are often coded in gene clusters, so called genomic islands. Therefore, the metagenome approach can also be applied to identify new genetic pathways for the directed synthesis of metabolites or enzymatic functions. Such functional genomic islands from uncultured microorganisms can be screened by isolating genetic material directly from original environmental samples (Fig. 1). The successful cloning and transformation of these sequences into a suitable host vector system for its expression and final characterization is a prerequisite for such screening approaches. One example in this respect is the commercialization of polyunsaturated fatty acids (PUFAs) from marine microorganisms. The most important PUFAs are eicosopentaenic acid (EPA) and docosahexaenic acid (DHA). PUFAs are of biotechnological interest because of their beneficial properties to human health and their importance in infant development [58]. The screening of microbial PUFA synthesis genes from metagenome libraries allows for the cloning of the responsible genes into suitable expression hosts. The cloning of a 38 kb gene cluster from Shewanella putrefaciens into E. coli and Synechococcus spp. resulted in a successful EPA production by these microorganisms [59, 60]. The potential knowledge of PUFA-related genes offers the possibility for construction of recombinant microbial cell factories suitable for an alternative production of EPA and DHA. The metagenome approach can also be used to automate the screening of genes from nature, either to look for new technical enzymes or for other specified activities. The screening of specific enzymatic activities requires the cloning and expression of a single gene. However, the production of distinct compounds, such as metabolites or antibiotics, is coded by a set of genes. The cloning of complex environmental DNA samples into a suitable host in combination with HTS methods is a good strategy for the screening and isolation of pathways for metabolic functions or the discovery of new bioactive natural compounds directly from the marine environment. One pioneer in this field is the company DIVERSA (San Diego, California). This company estimates that its gene expression (metagenome) libraries currently contain the complete genomes of over one million different microorganisms (http://www.diversa.com/). DIVERSA has developed a set of automated ultra HTS and enrichment strategies. The company uses two different screening strategies to discover novel biomolecules: (1) expression-based screening for biological activity and (2) sequence-based screening by identification of specific DNA sequences of interest. Recently, DIVERSA obtained the patent rights on a strategy claiming the construction and screening of expression libraries from nucleic acid directly isolated from the environment utilizing a fluorescence activated cell sorter (http://www.diversa.com/). This approach is based on a robotic screening system, which uses high-density microtiter

22

T. Schweder et al.

plates, capable of screening and characterizing of about one million clones per day. If a clone expresses an activity or contains a DNA sequence of interest, it is isolated for further analysis. 2.4 Functional Genome Analysis The function of the majority of genes within the sequenced marine genomes is not well understood. Furthermore, even if the complete set of genes of a microbial cell is available, it is mostly not known how these genes are regulated or how the proteins interact to express their functions. In order to assign potential functions to the genes of a genome, functional genome analysis techniques are used. These techniques include the expression profiling of the whole set of genes by using genomic DNA arrays and/or proteomics. These techniques are not only suitable for exploration of the functions of the proteins but also help to find new potential drug targets. Proteome and transcriptome analysis techniques have led to a shift from direct antimicrobial screening programs toward rational target-based strategies. Furthermore, these techniques allow the identification of essential genes for the synthesis of selected metabolites. 2.4.1 Proteome Analysis The proteome technique is mainly based on two-dimensional protein gel electrophoresis (2D-PAGE), used for protein separation, and MALDI-TOF mass spectrometry, applied for protein identification. Mandatory for the proteome analysis by mass spectrometry is the availability of the complete genome sequence of the organism of interest. The proteome of only few marine microorganisms has been investigated so far. Most of these proteome studies explore how marine bacteria adapt to alterations in their environmental conditions. One of the first proteome analyses was performed using a marine Vibrio [61, 62]. In these studies the response of the marine Vibrio sp. strain S14 to starvation for carbon, nitrogen, or phosphate and to simultaneous depletion of all these nutrients (multiple-nutrient starvation) was examined. Gross et al. (1994) investigated changes in the two-dimensional protein pattern of selected marine bacteria and fungi in response to high-pressure stress [63]. The determination of proteome signatures related to defined variations in the environmental conditions also allows the affiliation of unknown gene/protein functions into functional groups. Rabus et al. (2002) studied the proteome of the planctomycete Rhodopirellula baltica during growth with Nacetylglucosamine and glucose [64]. Analysis of the two-dimensional protein

Screening for New Metabolites from Marine Microorganisms

23

patterns revealed the presence of several protein spots, which were only detectable in soluble protein extracts of cells grown with N-acetylglucosamine. Lemus and Ngai (2000) examined alterations in the proteome of the Euprymna scolopes light organ in response to symbiotic Vibrio fischeri. 2D-PAGE identified changes in the soluble proteome of the symbiotic light organ induced by a specific response to the interaction with V. fischeri [65]. Lopez et al. (2002) suggested the application of proteomics for the identification of marine species by the analysis of species-specific peptides from randomly selected dominant protein spots [66]. 2.4.2 Transcriptome Analysis Proteome analysis is frequently accomplished by expression profiling with DNA arrays. However, genomic DNA arrays have so far been designed for only a few marine microorganisms. One of the first expression profiling studies with marine organisms using a genomic DNA array was done by Okamoto et al. (2003), who performed a genome-wide analysis of redoxregulated genes in a dinoflagellate [67]. In this study, the effects of sodium nitrite (a reactive nitrogen species generator) and paraquat (a producer of reactive oxygen species) on gene expression in the marine dinoflagellate species Pyrocystis lunula were investigated using genomic DNA microarrays. Beside the application of genomic DNA microarrays for gene expression profiling, the use of DNA chips, with probes for a selected number of gene sequences, for ecological surveys and the exploration of marine biodiversity has been suggested [68]. Wu et al. (2001) developed and evaluated functional DNA arrays with genes involved in nitrogen cycling from pure cultures and those cloned from marine sediments [70]. The authors could demonstrate that their model DNA array in principle reveals functional gene composition in natural microbial communities. However, the study also showed that more work is needed to improve the sensitivity and specificity of this method. The advantage of DNA chips in comparison to the FISH method would be that multiple numbers of probes can be applied in one hybridization experiment. This advantage of DNA chips is of special interest for the analysis of microbial communities with a complex biodiversity. Furthermore, in comparison to FISH this approach allows the application of multiple probes for a better control of false-positive and false-negative results. However, because of the large unknown amount of genetic sequences in an environmental sample, DNA chips are not yet widespread in marine microbial ecological applications [68]. For a successful application of DNA chips in this field, the discrimination of single mismatches is crucial [69].

24

T. Schweder et al.

2.5 Genome Sequencing and Identification of New Antimicrobial Compounds The sequencing of the genomes of microbial human pathogens has fundamentally changed the capabilities for antimicrobial drug screening [71] (Fig. 2). Most antimicrobial drugs used today are derivatives of structures originating from terrestrial microorganisms. However, these drugs have been applied for many years and a considerable number of pathogenic microorganisms have adapted to these structures by developing antibiotic resistances [72, 73]. Therefore, new lead structures allowing new mechanisms of

Fig. 2 The impact of genomics on drug development processes

Screening for New Metabolites from Marine Microorganisms

25

action, which are different from the established bioactive compounds from terrestrial microorganisms, are required. Most of the antimicrobial compounds currently on the market were screened based on whole cell antimicrobial screening programs. By application of new genome-driven techniques more directed, target-based approaches are possible [71]. These new screening strategies are directly coupled to potential drug targets, which have been identified by genome sequencing projects. Such antimicrobial targets are for example proteins that are essential for microbial growth or cell survival. Another important drug target are proteins that are involved in the pathogenicity of the pathogenic organism. In this respect, comparative genomics of pathogenic and non-pathogenic strains is a suitable approach for the identification of pathogenicity related proteins. For example, comparison of the genomes of uropathogenic E. coli strains with those of non-pathogenic E. coli and from other E. coli pathotypes revealed the existence of so-called pathogenicity islands [74–76]. A similar approach has been used to investigate the pathogenicity of the marine bacterium Vibrio vulnificus [77]. This halophilic marine bacterium is an etiologic agent of human mortality from seafood-borne infections. Genome sequencing and comparative analysis led to the identification of selected genomic regions that are typical for V. vulnificus and the human pathogen V. cholerae. The genomic information of this pathogenic bacterium can not only be applied for a monitoring of Vibrio infections but will eventually also lead to the identification of virulence factors of V. vulnificus. The sequencing of the genome of a microorganism that has been identified as a potent producer of bioactive compounds allows the identification of the gene clusters involved in the pathways for the production of these natural compounds. Myxobacteria like Myxococcus xanthus and Sorangium cellulosum have increasingly gained attention as producers of natural products with biological activity [78]. However, these myxobacteria are difficult to handle in large bioreactors for the biotechnological production of the metabolites of interests. Therefore, Müller and co-workers from the University of Saarbrücken are trying to identify biosynthetic gene clusters of these gram-negative bacteria by genome sequencing. In order to establish a biotechnological production of selected bioactive structures a suggestion is to clone the essential biosynthetic gene clusters of these myxobacteria into suitable hosts such as Pseudomonas or Streptomyces, which fit the demands of the myxobacterial genetics and biochemistry [78].

26

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3 Screening for New Metabolites 3.1 Alternative Cultivation Methods Less than 5% of the viable bacterial cells in marine samples ultimately grow under standard culture conditions [79, 80]. Typically, nutrient concentrations and cell numbers in the marine environment are three orders of magnitude lower than in common laboratory media [81]. Based on this fact, Button et al. (1993) developed a new approach for the isolation of marine bacteria [82]. This group simulated the normal substrate concentrations but also the cell density limited environmental conditions and was able to isolate typical marine bacteria. Rappe et al. [81] have improved the technique of Button et al. [82] and applied microarrays from the cell cultures coupled with FISH. A high throughput procedure allowing the cultivation of the abundant group of the ubiquitous SAR11 marine bacterioplankton clade was used. The combination of the HTS and molecular analysis techniques was helpful in increasing the rate at which cultures could be obtained and identified [81]. Fresh Oregon coast seawater samples were diluted so that each well of a microtiter plate was inoculated on average with about 22 microbial cells. The medium in the microtiter wells was sterile Oregon coast water supplemented with phosphate, ammonium, and defined mixtures of organic carbon compounds. This approach allowed isolation of 18 bacterial cultures, which were not cultivable under standard cultivation conditions and which had resisted any cultivation efforts in the past. Rappe et al. [81] thus confirmed the suitability of diluting natural microbial communities in very low nutrient media for the isolation of new marine microorganisms. Consequently, high similarity to the original environmental conditions of the samples is helpful for the isolation of novel microbial species. Low nutrient conditions and the establishment of the selection of single cells for cultivation were also successfully applied for the isolation of novel halophiles from Red Sea brine [83]. Furthermore, it has been reported that a combination of FISH and microautoradiography supports the determination of physiological activities of microorganisms without cultivation [84]. In this approach a radioactively labeled substrate was used for the incubation of a complex microbial sample. Cells metabolizing the radioactive-labeled substrate were finally detected by FISH. By this technique the strains are not only taxonomically affiliated but it is also possible to gain information concerning their physiological activity and the utilization of distinct substrates. There is a challenge of developing further new culture techniques that incorporate an understanding of the special environmental conditions of marine organisms in their natural habitats. This includes variations of dissolved

Screening for New Metabolites from Marine Microorganisms

27

organic matter, trace elements, or surfaces. In most cases essential interactions with other organisms of the marine biotope have to be considered. The growth and development of selected microorganisms could directly or indirectly depend on the association with other bacteria or algae in the marine environment. Therefore, the consideration of microbial consortia rather than purified cultures could be a further suitable strategy for screening for new microbial activities. Furthermore, most cultivation attempts are performed either in liquid media or on agar plates as batch cultures. Special cultivation conditions, such as permeable solid substrates that imitate sediments or continuous cultivations in flow reactors, are rarely applied. Another interesting aspect is the potential addition of growth-supporting signal molecules. Bruns et al. (2002) used signaling factors like the autoinducer acyl homoserine lactone or cyclic AMP to support the culturing of marine microorganisms [85]. It is thought that microorganisms only synthesize their bioactive compounds under distinct environmental conditions. Thus, beside the cultivation of these microorganisms the identification of the most suitable growth conditions for a maximal production of secondary metabolites also has to be considered. The influence of culture conditions on secondary metabolite production has for example been demonstrated for cyclic and linear lipopeptides of Cyanobacteria [86]. The competition of different microorganisms for limited natural resources is also supposed to support the synthesis of antimicrobial compounds. In the marine bacterium Streptomyces tenjimariensis production of the antibiotic istamycin is induced by co-culturing with other marine bacteria [87]. Such antibiotics frequently inhibit the growth of competitors and thus play an ecological role in the suppression of competitive microorganisms. 3.2 Preparation of Materials for Screening The screening results depend on the quality of screening material, collection and storage of organisms, cultivation, extraction, storage of extracts, and preparation of test samples. A directed (preselected) screening offers better chances of finding interesting metabolites than an undirected (“blind”) screening. Such a directed screening could be based on ecological observations (peculiarities like interactions, special environmental conditions), on traditional experiences (use in ethnomedicine), or search in novel organisms. Mode and solvent of extraction determine which substances are extracted [88, 89]. Solid phase extraction is a suitable method for automated sample preparation [90]. Extracts make special demands on sample preparation and screening systems because they are usually complex mixtures of compounds. The concentration of active compounds in crude extracts is often very low (1 : 1000). One extract could contain compounds with opposite effects or compounds with

28

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low effect, which are active only in combination. False-positive results could be found. Further problems could be the lack of solubility in physiological solvents, the coloring of an extract (difficulties in spectroscopic measurements), or the non-tolerability of crude extracts to some bioassays. 3.3 Chemical and Physicochemical Screening Chemical and physicochemical screening is the search for new chemical structures regardless of their biological activities. The first step is the separation of compounds from a complex extract by chromatographic methods. In a second step, the chemical reactivity or physicochemical properties of the separated compounds are analyzed by spectroscopic methods (UV/VIS, MS, NMR) or by detection with special detection reagents in the TLC [91]. The development of HPLC-DAD-MS systems allows the specific detection of single components in a complex mixture (e.g., an extract), regardless of the background of other metabolites. Comparison with data bases allows the identification of known structures (dereplication). Finding of unknown peaks in a HPLC or TLC chromatogram stimulates the isolation of the compound giving this peak by chromatographic methods and allows structure elucidation of the isolated substance by spectroscopic methods like nuclear magnetic resonance spectroscopy (NMR) or mass spectroscopy (MS). Taking into account the growth of (marine) microorganisms in microtiter plates, the screening for secondary metabolites could also be done on this miniaturized scale [92]. The new metabolites found by chemical screening still have to be tested for their biological activities. Combination of HPLC-separation, bioassays, and on-line spectroscopy miniaturizes and accelerates the identification of bioactive metabolites in a complex matrix [93]. 3.4 Biological Screening During biological screening test samples (extracts, fractions, pure compounds, and compound libraries) are screened for their bioactivities in vitro and/or in vivo. In the case of extracts, active metabolites could be isolated by bioactivity-guided isolation processes. The finding of structurally known compounds (dereplication) in active extracts is possible. Biological screening methods have to be of relevance for the objective (e.g., for the therapeutic aim), highly sensitive, selective (insensitive to inactive compounds and to ubiquitous compounds), and reproducible. They should be characterized by high information content and reasonable costs [94]. In vitro tests could be done on a molecular or on a cellular level. The information content will increase from molecular to cellular level, but the

Screening for New Metabolites from Marine Microorganisms

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throughput is higher in molecular test systems (question of quantity vs. quality). Assays that require careful interpretation but provide a lot of information per assay are ideal for marine natural products research [88]. Tests on the molecular level are based, e.g., on receptor systems (identification of those compounds which bind to one receptor) or on enzyme systems (enzyme-catalyzed turnover rates). Tests on the genome, transcriptome, or proteome level will become more and more important. Targets of high pharmacological relevance are G-protein coupled receptors, tyrosine kinase receptors, nuclear hormone receptors, ion channels, proteases, kinases, phosphatases, and transporter molecules. The detection of a reaction on the molecular level could be done by biochemical assays (e.g., spectrophotometric measurement of the product of an enzymatic reaction), ligand binding assays (readout by labeling with a tracer) or functional assays (reporter gene assays quantifying the expression level of a specific reporter gene product, second messenger assays, two hybrid assays for measuring protein–protein interactions). Fluorescence-based assay technologies, isotopic labeling, colorimetry, and chemoluminescence are very often used as detection methods [95–97]. Cell-based assays are more complex and more physiologically relevant than tests on the molecular level. On the other hand they are still labor intensive and more difficult to validate than molecular assays [98]. Permanent cell lines have many advantages in handling compared to primary cell cultures. Examples for cellular assays are: (1) vitality and proliferation studies with tumor cells or other permanent cell lines for the detection of antitumor effects or undesired cytotoxic effects, and (2) growth studies with bacterial or fungal cells (diffusion assay or dilution assay) for detection of antibiotic effects or morphological and biochemical investigations of special cell types, e.g., endothelial cells, muscle cells, or keratinocytes. The detection could be done by microscopic investigation, counting of cell number, measurement of electrophysiological properties, determination of metabolic capacities or membrane integrity (dye uptake assay), uptake of precursors into DNA (proliferation studies), or several biochemical parameters (protein content and so on). Most marine samples have only been tested in a limited number of assays. The true drug potential of many of these compounds may therefore not yet be fully realized. Papers about new chemical structures of marine compounds give only initial biotesting data. Their significance is often difficult to determine. Papers with systematic screening results obtained from extracts are rare. The goal of highlighting compounds that are likely to become clinical candidates is also complicated by the fact that pharmaceutical companies are naturally reluctant to talk about compounds in the early stages of development [88]. The results of detailed pharmacological and biochemical studies could therefore be found only for selected compounds. A predominant portion of bioactive marine metabolites (from marine bacteria and fungi, but also from invertebrates) is occupied by antitumor/cytotoxic compounds. Other activities are antibacterial, antiviral, anti-inflamma-

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tory, and enzyme-inhibiting [99]. Some toxic metabolites could be useful as molecular tools. 3.5 High-Throughput Screening, Automation, Data Management The use of high-throughput screening (HTS) methods can improve the efficiency of biological tests [95]. HTS is characterized by very high numbers of test samples (more than 100 000 per day in UHTS), automated processes, single-time-point measurement instead of kinetic measurements, development of miniaturized and paralleled techniques (96-, 384- or 1536-well microtiter plates), and more sensitive detection technologies with fast read time [96, 100]. Presently HTS or UHTS are designed to handle large libraries of pure compounds, e.g., obtained by combinatorial chemistry [101]. They mainly use isolated targets. The development of new HTS methods for cell-based screening is in progress. Because of their higher complexity they could simultaneously deliver information on multiple parameters, provide higher quality data, and give better information about function of bioactive principles. Reporter systems are introduced into the cells, which are based on the expression of genes encoding proteins such as β-galactosidase, luciferase, or alkaline phosphatase. An alternative to the use of expensive mammalian cells is the expression of a selected physiological process in microorganisms such as Saccharomyces cerevisiae. Novel methods of single molecule detection, e.g., fluorescence correlation spectroscopy (FCS) will improve the processes [95, 102]. High-content screening (HCS) becomes more and more important. This means that a smaller number of high-qualitative compounds (higher purity) is screened with higher performance (high information content). HTS is performed mainly by the industry and seldom by academic groups. It focuses on the active principle in a distinct bioassay. The screens are usually run only for a limited length of time. Due to the above mentioned problems with extracts, HTS is not suitable for these samples and crude extracts derived from natural sources therefore only play a minor role in HTS [95]. A bioassayguided fractionation of an extract could need longer time than the screening of the pure chemical library [88]. A way to overcome the problems of biogenic samples in HTS is the synergistic use of natural product chemistry and combinatorial chemistry. By bringing together the structural value and complex molecular shape of natural products isolated from a natural source with rational synthetic strategies of combinatorial chemistry and biochemistry, the success of screening processes (the chance to find new lead structures) could be significantly improved [103]. Thus one could randomly isolate pure compounds on the basis of interesting chemical structures, produce substance libraries and then screen these libraries [88].

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In every case, the results of HTS represent only the first step in a series of experiments; valid cell culture experiments and animal assays are necessary. ADMET properties (adsorption, distribution, metabolism, excretion, and toxicology of a bioactive metabolite) have to be considered. Only compounds with positive results in primary HTS and in secondary assays could be declared as a “hit”. The hits have also to be structurally defined. Derivatization of hits by chemical or biochemical methods could produce a “lead”, a series of hits for which the structure-activity relationship is shown and activity demonstrated in vitro and in vivo. A new development in the field of HTS is represented by the High Throughput Pharmacological System™ (HTPS), which was patented by Axiom Biotechnologies (San Diego, CA) [104]. This is a fluidics- based platform that uses viable cells and test compounds to identify active compounds. This approach allows for a very fast estimation of the potency of the compounds and a determination of their specificity. HTPS can be understood as an instrument for automated programming of complex pharmacological cell treatment protocols. Edwards et al. (2001) [105] have coupled HTPS to a flow cytometer using a plug flow coupling valve technology. Flow cytometry is performed with fluorescent probes and allows an optical measurement of physiological parameters of individual cells or cellular macromolecules at a high rate [106]. HTPS flow cytometry facilitates a high-throughput multifactorial screening of large compound libraries to increase the efficiency with which novel bioresponse modifying drugs, such as from marine microorganisms, may be identified and characterized [105]. In this approach a large compound library in microtiter plates is sequentially combined with cells and finally delivered to a flow cytometer for multiparametric analysis. 3.6 Metabolome Analysis Techniques The metabolome is the final product of proteome activity including the total assembly of low molecular weight molecules in a cell. Its composition is determined not only by the genetic information encoded in the genes, but also by a particular physiological and developmental state as well as by environmental factors. The main technologies for metabolome analysis are MS and NMR spectrometry, often combined with chromatographic methods. These require minute amounts of sample and will accommodate individual components of highly varying chemical structures and physical properties. Metabolome analysis is expected to become a valuable tool in the search for new metabolites from marine microorganisms and in investigation of their biological effects on the metabolome of other cells. Two of the methods of choice for the evaluation of metabolic constituents of cells are the hyphenated techniques HPLC-NMR and HPLC-MS. These methods can be used for the identification of several individual compo-

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nents present in mixtures. Rapid identification is possible by comparison to database entries of the NMR and MS patterns of compounds of interest [107, 108]. By intelligent coupling of metabolome analysis with cellular test systems it seems possible to detect the effects (or side effects) of drug candidates at a very early stage of the drug development process and so to reduce the late stage failure of compounds in the pipeline. 3.7 Examples for Metabolites from Marine Microorganisms Some excellent reviews about bioactive metabolites from marine microorganisms are given by Fenical and Jensen [79, 109, 110]. Selected examples will be outlined in this overview. Antitumor agents Test samples are screened in cell-based assays against a panel of different human tumor cell lines. The NCI uses about 60 cell lines representing nine of the most important tumor types. In the NCI screening the percentage of significant active cytotoxic samples (IC 50 < 4 µg/mL) among marine organisms was higher (2%) than that among plants (< 1%) [111]. After primary screens numerous sophisticated molecular and biochemical screens that target specific cellular aspects of cancer growth and dissemination should follow [110]. A highlighted metabolite is salinosporamide A from a marine bacterium of the new genus Salinospora, a group of obligate marine actinomycetes widely distributed in marine sediments (Fig. 3). These bacteria possess a γ lactam-β-lactone bicyclic ring structure with clasto-lactacystin-β-lactone with unique functionalization displaying potent in vitro cytotoxicity (mean IC 50 value less than 10 nM). In more sophisticated tests it could be shown that salinosporamide A inhibits the 20S subunit of proteasomes [112]. Due to its essential role in cellular physiology the proteasome represents a very attractive target for new drugs. Other examples for potential antitumor drugs from marine microorganisms are the alkaloid alteramide (Fig. 3) from a marine bacterium (Alteromonas) isolated from the sponge Halichondria okadai [113], the diketopiperazine dimer asperazine (Fig. 3) from a strain of Aspergillus niger obtained from the sponge Hyrtios sp. [114], the macrolide halichomycin (Fig. 4) from Streptomyces hygroscopicus, isolated from the gastrointestinal tract of the marine fish Halichoeres bleekeri [115], the depsipeptide thiocoraline (Fig. 4) from Micromonospora marina [116], the leptosins, diketopiperazine dimers from the obligate marine fungus Leptosphaeria sp. [117], and the neomangicols, partly halogenated sesterterpenes isolated from the mycelial extract of a wood-inhabiting marine fungus (Fig. 5) [118].

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Fig. 3 Antitumor compounds from marine microorganisms (I)

Fig. 4 Antitumor compounds from marine microorganisms (II)

The IC 50 values of these compounds against various cell lines range from 0.1 to less than 10 µg/mL. More significantly, leptosins A and C displayed potent in vivo activity in a Sarcoma-180 ascites tumor model in mice [110].

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Fig. 5 Antitumor compounds from marine microorganisms (III)

Fig. 6 Antitumor compounds from marine microorganisms (IV)

Cytotoxic metabolites from cyanobacteria are cryptophycin from a Nostoc sp. [119] and curacin A from a specimen of Lyngbya majuscula from Curac¸ao, which inhibits microtubule assembly by binding at the colchicin site (Fig. 6). Curacin A is only active in vitro [120]. The potent cytotoxin dolastatin-10, primarily found in the sea hare Dolabella auricularia, has recently been isolated from a cyanobacterium (Sym-

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Fig. 7 Antitumor compounds from marine microorganisms (V)

ploca sp., [121]). The lyngbyastatins are cytotoxic depsipeptides from Lyngbya majuscula (Fig. 7) [122]. Antibiotics Agar diffusion assays using different microbial strains (including multiresistant strains) represent the most important screening method for detection of antibacterial and antifungal effects of extracts and pure compounds. Active test samples are further evaluated by dilution assays to determine the minimal inhibitory concentration and by cytotoxicity assays to exclude undesired effects against mammalian cells. Furthermore, tests for determination of target and mode of action (see Sect. 4) and in vivo assays are necessary. Examples for new antibacterial agents from marine microorganisms are the massetolides A–H (Fig. 8), cyclic depsipeptides from two Pseudomonas strains obtained from a marine alga and a tube worm [123], marinone and debromomarinone from a marine-sediment derived actinomycete [124], microsphaeropsisin from a fungus (Microsphaeriopsis sp.) isolated from a sponge [125], ascochitin and ascochital from Kirschsteiniothelia maritima

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Fig. 8 Antibiotic compounds from marine microorganisms (I)

Fig. 9 Antibiotic compounds from marine microorganisms (II)

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Fig. 10 Antifungal compounds from marine microorganisms

isolated from wood [126], and corollosporin from the fungus Corollospora maritima obtained from wood nearby Helgoland, Germany (Fig. 9) [127]. Antifungal activities are exhibited by lipodepsipeptides (LL-15 G256) from the fungus Hypoxylon oceanicum (Fig. 10). They inhibit the fungal cell wall synthesis [128]. Antiviral agents In vitro tests for antiviral activity are carried out in cellular test systems such as the influenza virus/MDCK cell system (Martin-Darby Canine Kidney cells) or herpes simplex virus/VERO cells (kidney cells from green macaque). Cells are damaged by virus infection and antiviral effects can be shown by improved cell viability or membrane integrity. Other test methods are based on virus specific enzymes. Examples for antiviral agents from marine microorganisms are the macrolactins A–F, macrolides from a Gram-positive deep-sea bacterium from a sediment-sample obtained from the California coast [129], the caprolactins A and B containing a cyclized lysine moiety also from a Gram-positive deep-sea bacterium from a sediment sample [130], the halovirs A–C, cyclic peptides similar to peptaibols from a marine fungus (Scytalidium sp., [110]), and sulfated polysaccharides from a marine Pseudomonas sp. (Fig. 11) [131]. The named compounds inhibit herpes simplex viruses; in addition macrolactin A is effective against HIV.

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Fig. 11 Antiviral compounds from marine microorganisms

Anti-Inflammatory agents Anti-inflammatory effects can be shown by the inhibition of enzymes involved in the inflammation process (cyclooxygenases, lipoxygenases, some proteases), on production and release of inflammatory cytokines, on blood coagulation, histamine release, or in animal assays (phorbol ester induced ear edema in mice and other). Examples for anti-inflammatory metabolites from marine microorganisms are the salinamides A and B, depsipeptides from an actinomycete isolated from the surface of the jelly fish Cassiopeia xamancha [132]; thiotropocin, a sulfur-containing macrolide from a Caulobacter sp. [133]; and the phomactins A–G, sesquiterpenes from a Phoma sp. obtained from a crab shell (Fig. 12) [134]. The salinamides inhibit inflammation in a mouse ear edema assay by > 80% at a dose of 50 µg/ear [132], phomactin D inhibits PAF (plateletactivating factor)-induced platelet aggregation with an IC50 value of 0.8 µM [134], and thiotropocin depresses histamine release and hemolysis of rabbit erythrocytes [133].

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Fig. 12 Anti-inflammatory compounds from marine microorganisms

Agents with other indications Enzyme inhibitors are of interest for treatment of several diseases, e.g., cardiovascular or kidney diseases. They are the focus of many in vitro screening programs. A novel endothelin-converting enzyme inhibitor (compound B-90063) with 4-pyridone and oxazole skeletons was discovered to be produced by a new marine species of the genus Blastobacter, isolated from seawater (Fig. 13) [135]. Endothelin is produced by endothelial cells and participates in the regulation of blood pressure. Inhibitors are therefore of interest for treatment of cardiovascular diseases. The polyketide obionin from the obligate marine fungus Leptosphaeria obiones, isolated from the coastal march grass Spartina alterniflora, inhibits ligand binding to a dopamine-selective receptor with an IC 50 of 2.5 µg/mL (Fig. 13) [136]. Komodoquinone A from a marine Streptomyces sp. [137] and epolactaene, a polyene from a Penicillium sp. [138], induce the differentiation of neuronal cells and could be of interest for regeneration of nerve cells (Fig. 14). The semiplenamides A–G are fatty acid amides from a Papua New Guinea collection of the marine cyanobacterium Lyngbya majuscule which influence the endogenic cannabinoid system of humans (Fig. 14) [139].

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Fig. 13 Compounds with other indications from marine microorganisms (I)

Fig. 14 Compounds with other indications from marine microorganisms (II)

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4 Application of Proteomics for Target Analyses of Antibacterial Compounds Proteome analysis is a suitable experimental tool for evaluating the mode of action of drugs [140, 141]. In the case of known drugs proteome analysis with suitable microbial model cells may confirm and complement the results of biochemical and other target analysis assays. Comparisons to antibiotics with known targets can be used to confirm the observed correlations. In the case of new drugs, the target of which is unknown, proteome signatures give the first indications of their mode of action. By comparing treated cells to untreated controls, the influence of the drug on protein patterns of the model cells can be followed. By this approach it was shown that the antibacterial compounds ascochitin, ascochital, and corollosporin (Fig. 9), isolated from marine fungi, induce protein stress in Bacillus subtilis. By comparison to protein signatures of known antibiotics an influence on translation by these antimicrobial compounds could be proposed [142].

5 Influence of Cultivation Conditions on Metabolite Production Genomic analysis has shown that most fungi or bacteria that produce secondary metabolites have the genetic potential for several biosynthetic pathways and therefore to generate more than one compound. By alteration of cultivation parameters it is possible to increase the number of secondary metabolites from one microbial source (one strain – many compounds, OSMAC) [143]. Chemical and biological screening methods are necessary to record the whole spectrum of metabolites. This approach offers a good alternative to industrial HTS, where the detection of additional compounds in extracts that might be of interest in other bioassays is impossible. By a systematic approach, the group of Zeeck [143] was able to isolate more than 100 compounds belonging to more than 25 different structural classes from only six different terrestrial microorganisms. It is supposed that marine microorganisms show similar abilities. Functional genome analysis will help to identify the cultivation conditions that activate silent gene clusters responsible for the biosynthesis of bioactive metabolites and the molecular mechanism triggering this change in expression profile.

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6 Outlook Exploitation of the potential of marine microorganisms as producers of bioactive metabolites is just beginning. For the realization of this potential, it is necessary that the full extent of biological and chemical diversity is examined by appropriate methods. Until now traditional screening methods limited to selected disease areas have mostly been used. Methods of functional genome analysis including transcriptome, proteome, and metabolome analysis will lead to a better understanding of disease mechanisms and the identification of new drug targets. Improved screening methods and further development of the necessary technical equipment will allow immense progress in the drug development process. They will also promote the cultivation of until now uncultivable marine microorganisms and investigation of their genetic potential. It is expected that such novel and obligate marine organisms contain more new metabolites than easily accessible facultative marine microorganisms. The exploration of the genome structure and physiology of uncultivable microorganisms by environmental genomics will lead to the discovery of new biotechnologically relevant enzymes and genes that code for bioactive metabolites. By improving the HTS of metagenome databases complete genomic coverage of special marine habitats will become possible. The revolutionary development in the field of functional genomics during the last 10 years supports this assumption. Nevertheless, it should be clear that years of development and a lot of money are required before utility of new marine natural compounds in medicine and biotechnology is fully accomplished.

References 1. Lindequist U, Schweder T (2001) In: Rehm HJ, Reed G (eds) Biotechnology, vol 10: Special processes. Wiley VCH, pp 441–484 2. Staley, J (2002) Biodiversity of microbial life. Wiley-Liss, New York 3. Smith DR, Doucette-Stamm LA, Deloughery C, Lee H, Dubois J, Aldredge T, Bashirzadeh R, Blakely D, Cook R, Gilbert K, Harrison D, Hoang L, Keagle P, Lumm W, Pothier B, Qiu D, Spadafora R, Vicaire R, Wang Y, Wierzbowski J, Gibson R, Jiwani N, Caruso A, Bush D, Reeve JN, et al. (1997) J Bacteriol 179:7135–55 4. Nakamura Y, Kaneko T, Hirosawa M, Miyajima N, Tabata S (1998) Nucleic Acids Res 26:63–67 5. Deckert G, Warren PV, Gaasterland T, Young WG, Lenox AL, Graham DE, Overbeek R, Snead MA, Keller M, Aujay M, Huber R, Feldman RA, Short JM, Olsen GJ, Swanson RV (1998) Nature 392:353–358

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6. Kawarabayasi Y, Hino Y, Horikawa H, Yamazaki S, Haikawa Y, Jin-no K, Takahashi M, Sekine M, Baba S, Ankai A, Kosugi H, Hosoyama A, Fukui S, Nagai Y, Nishijima K, Nakazawa H, Takamiya M, Masuda S, Funahashi T, Tanaka T, Kudoh Y, Yamazaki J, Kushida N, Oguchi A, Kikuchi H, et al. (1999) DNA Research 6:83–101 7. Golyshin PN, Martins Dos Santos VA, Kaiser O, Ferrer M, Sabirova YS, Lunsdorf H, Chernikova TN, Golyshina OV, Yakimov MM, Puhler A, Timmis KN (2003) J Biotechnol 106:215–220 8. Kaneko T, Nakamura Y, Wolk CP, Kuritz T, Sasamoto S, Watanabe A, Iriguchi M, Ishikawa A, Kawashima K, Kimura T, Kishida Y, Kohara M, Matsumoto M, Matsuno A, Muraki A, Nakazaki N, Shimpo S, Sugimoto M, Takazawa M, Yamada M, Yasuda M, Tabata S (2001) DNA Res 8:205–13; 227–253 9. http://wit.integratedgenomics.com/GOLD/index.cgi?want=Prokaryotic+Ongoing+ Genomes 10. Eisen JA, Nelson KE, Paulsen IT, Heidelberg JF, Wu M, Dodson RJ, Deboy R, Gwinn ML, Nelson WC, Haft DH, Hickey EK, Peterson JD, Durkin AS, Kolonay JL, Yang F, Holt I, Umayam LA, Mason T, Brenner M, Shea TP, Parksey D, Nierman WC, Feldblyum TV, Hansen CL, Craven MB, Radune D, Vamathevan J, Khouri H, White O, Gruber TM, Ketchum KA, Venter JC, Tettelin H, Bryant DA, Fraser CM (2002) Proc Natl Acad Sci USA 99:9509–9514 11. http://www.tigr.org/tdb/mdb/mdbinprogress.html 12. Saunders NFW, Thomas T, Curmi PMG, Mattick JS, Kuczek E, Slade R, Davis J, Franzmann PD, Boone D, Rusterholtz K, Feldman K, Gates C, Bench S, Sowers K, Kadner K, Aerts A, Dehal P, Detter C, Glavina T, Lucas S, Richardson P, Larimer F, Hauser L, Land M, Cavicchioli R (2003) Genome Res 13:1580–1588 13. Bult CJ, White O, Olsen GJ, Zhou L, Fleischmann RD, Sutton GG, Blake JA, FitzGerald LM, Clayton RA, Gocayne JD, Kerlavage AR, Dougherty BA, Tomb JF, Adams MD, Reich CI, Overbeek R, Kirkness EF, Weinstock KG, Merrick JM, Glodek A, Scott JL, Geoghagen NS, Venter JC (1996) Science 273:1058–1073 14. Slesarev AI, Mezhevaya KV, Makarova KS, Polushin NN, Shcherbinina OV, Shakhova VV, Belova GI, Aravind L, Natale DA, Rogozin IB, Tatusov RL, Wolf YI, Stetter KO, Malykh AG, Koonin EV, Kozyavkin SA (2002) Proc Natl Acad Sci USA 99:4644–4649 15. Waters E, Hohn MJ, Ahel I, Graham DE, Adams MD, Barnstead M, Beeson KY, Bibbs L, Bolanos R, Keller M, Kretz K, Lin X, Mathur E, Ni J, Podar M, Richardson T, Sutton GG, Simon M, Soll D, Stetter KO, Short JM, Noordewier M (2003) Proc Natl Acad Sci USA 100:12 984–12 988 16. Takami H, Takaki Y, Uchiyama I (2002) Nucleic Acids Res 30:3927–3935 17. Dufresne A, Salanoubat M, Partensky F, Artiguenave F, Axmann IM, Barbe V, Duprat S, Galperin MY, Koonin EV, Le Gall F, Makarova KS, Ostrowski M, Oztas S, Robert C, Rogozin IB, Scanlan DJ, Tandeau de Marsac N, Weissenbach J, Wincker P, Wolf YI, Hess WR (2003) Proc Natl Acad Sci USA 100:10 020–10 025 18. Rocap G, Larimer FW, Lamerdin J, Malfatti S, Chain P, Ahlgren NA, Arellano A, Coleman M, Hauser L, Hess WR, Johnson ZI, Land M, Lindell D, Post AF, Regala W, Shah M, Shaw SL, Steglich C, Sullivan MB, Ting CS, Tolonen A, Webb EA, Zinser ER, Chisholm SW (2003) Nature 424:1042 19. Fitz-Gibbon ST, Ladner H, Kim UJ, Stetter KO, Simon MI, Miller JH (2002) Proc Natl Acad Sci USA 99:984–989 20. Cohen GN, Barbe V, Flament D, Galperin M, Heilig R, Lecompte O, Poch O, Prieur D, Querellou J, Ripp R, Thierry JC, Van der Oost J, Weissenbach J, Zivanovic Y, Forterre P (2003) Mol Microbiol 47:1495–1512

44

T. Schweder et al.

21. Robb FT, Maeder DL, Brown JR, DiRuggiero J, Stump MD, Yeh RK, Weiss RB, Dunn DM (2001) Methods Enzymol 330:134–157 22. Kawarabayasi Y, Sawada M, Horikawa H, Haikawa Y, Hino Y, Yamamoto S, Sekine M, Baba S, Kosugi H, Hosoyama A, Nagai Y, Sakai M, Ogura K, Otsuka R, Nakazawa H, Takamiya M, Ohfuku Y, Funahashi T, Tanaka T, Kudoh Y, Yamazaki J, Kushida N, Oguchi A, Aoki K, Kikuchi H (1998) DNA Research 5:55–76 23. http://www.spaceref.com/news/viewpr.html?pid=6101 24. Glockner FO, Kube M, Bauer M, Teeling H, Lombardot T, Ludwig W, Gade D, Beck A, Borzym K, Heitmann K, Rabus R, Schlesner H, Amann R, Reinhardt R (2003) Proc Natl Acad Sci USA 100:8298–8303 25. Kawarabayasi Y, Hino Y, Horikawa H, Jin-no K, Takahashi M, Sekine M, Baba S, Ankai A, Kosugi H, Hosoyama A, Fukui S, Nagai Y, Nishijima K, Otsuka R, Nakazawa H, Takamiya M, Kato Y, Yoshizawa T, Tanaka T, Kudoh Y, Yamazaki J, Kushida N, Oguchi A, Aoki K, Masuda S, Yanagii M, Nishimura M, Yamagishi A, Oshima T, Kikuchi H (2001) DNA Research 31:123–140 26. Palenik B, Brahamsha B, Larimer FW, Land M, Hauser L, Chain P, Lamerdin J, Regala W, Allen EE, McCarren J, Paulsen I, Dufresne A, Partensky F, Webb EA, Waterbury J (2003) Nature 424:1037–1042 27. Kaneko T, Sato S, Kotani H, Tanaka A, Asamizu E, Nakamura Y, Miyajima N, Hirosawa M, Sugiura M, Sasamoto S, Kimura T, Hosouchi T, Matsuno A, Muraki A, Nakazaki N, Naruo K, Okumura S, Shimpo S, Takeuchi C, Wada T, Watanabe A, Yamada M, Yasuda M, Tabata S (1996) DNA Research 3:109–136 28. Nakamura Y, Kaneko T, Sato S, Ikeuchi M, Katoh H, Sasamoto S, Watanabe A, Iriguchi M, Kawashima K, Kimura T, Kishida Y, Kiyokawa C, Kohara M, Matsumoto M, Matsuno A, Nakazaki N, Shimpo S, Sugimoto M, Takeuchi C, Yamada M, Tabata S (2002) DNA Research 9:123–130 29. Nelson KE, Clayton RA, Gill SR, Gwinn ML, Dodson RJ, Haft DH, Hickey EK, Peterson JD, Nelson WC, Ketchum KA, McDonald L, Utterback TR, Malek JA, Linher KD, Garrett MM, Stewart AM, Cotton MD, Pratt MS, Phillips CA, Richardson D, Heidelberg J, Sutton GG, Fleischmann RD, Eisen JA, Fraser CM, et al. (1999) Nature 399:323–329 30. http://ergo.integratedgenomics.com/Genomes/VFI/index.html 31. Chung-Yung C, Keh-Ming W, Yo-Cheng C, Chuan-Hsiung C, Hui-Chi T, Tsai-Lien L, Yen-Ming L, Hsiang-Ju C, Bo-Ting AS, Jian-Chiuan L, Teh-Li S, Chung-Ping S, Chung-Te L, Lien-I H, Shih-Feng T (2003) Genome Res 13:2577–2587 32. http://wit.integratedgenomics.com/GOLD/index.cgi?want=Prokaryotic+Ongoing+ Genomes 33. Schüler D, personal communication 34. http://www.sanger.ac.uk/Projects/M_marinum/ 35. Danchin A, personal communication 36. Dufresne A, Salanoubat M, Partensky F, Artiguenave F, Axmann IM, Barbe V, Duprat S, Galperin MY, Koonin EV, Le Gall F, Makarova KS, Ostrowski M, Oztas S, Robert C, Rogozin IB, Scanlan DJ, Tandeau de Marsac N, Weissenbach J, Wincker P, Wolf YI, Hess WR (2003) Proc Natl Acad Sci USA 100:10 020–10 025 37. Rocap G, Larimer FW, Lamerdin J, Malfatti S, Chain P, Ahlgren NA, Arellano A, Coleman M, Hauser L, Hess WR, Johnson ZI, Land M, Lindell D, Post AF, Regala W, Shah M, Shaw SL, Steglich C, Sullivan MB, Ting CS, Tolonen A, Webb EA, Zinser ER, Chisholm SW (2003) Nature 424:1042–1047 38. Partensky F, Hess WR, Vaulot D (1999) Microbiol Mol Biol Rev 63:106–127

Screening for New Metabolites from Marine Microorganisms

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39. Glöckner FO, Kube M, Bauer M, Teeling H, Lombardot T, Ludwig W, Gade D, Beck A, Borzym K, Heitmann K, Rabus R, Schlesner H, Amann R, Reinhardt R (2003) Proc Natl Acad Sci USA 100:8298–8303 40. Distel DL, Lane DJ, Olsen GJ, Giovannoni SJ, Pace B, Pace NR, Stahl DA, Felbeck H (1988) J Bacteriol 170:2506–2510 41. Paul JH, Sullivan MB, Segall AM, Rohwer F (2002) Comp Biochem Physiol Part B 133:464–476 42. Liu J, Dehbi M, Moeck G, Arhin F, Bauda P, Bergeron D, Callejo M, Ferretti V, Ha N, Kwan T, McCarty J, Srikumar R, Williams D, Wu JJ, Gros P, Pelletier J, DuBow M (2004) Nat Biotechnol 22:185–191 43. Olsen GJ, Woese CR (1993) FASEB J 7:113–123 44. Ludwig W, Strunk O, Klugbauer S, Klugbauer N, Weizenegger M, Neumaier J, Bachleitner M, Schleifer KH (1998) Electrophoresis 19:554–568 45. DeLong EF (2001) Curr Opin Microbiol 4:290–295 46. Hugenholtz P, Goebel BM, Pace NR (1998) J Bacteriol 180:4765–4774 47. Amann R, Fuchs BM, Behrens S (2001) Curr Opin Biotechnol 12:231–236 48. DeLong EF, Schleper C, Feldman R, Swanson RV (1999) Biol Bull 196:363–365 49. Handelsman J, Rondon MR, Brady SF, Clardy J, Goodman RM (1998) Chem Biol 5:245–249 50. Rodriguez-Valera F (2002) Environ Microbiol 4:628–633 51. Stein JL, Marsh TL, Wu KY, Shizuya H, DeLong EF (1996) J Bacteriol 178:591–599 52. Beja O, Suzuki MT, Koonin EV, Aravind L, Hadd A, Nguyen LP, Villacorta R, Amjadi M, Garrigues C, Jovanovich SB, Feldman RA, DeLong EF (2000) Environ Microbiol 2:516–29 53. Beja O, Aravind L, Koonin EV, Suzuki MT, Hadd A, Nguyen LP, Jovanovich SB, Gates CM, Feldman RA, Spudich JL, Spudich EN, DeLong EF (2000) Science 289:1902– 1906 54. Shizuya H, Birren B, Kim UJ, Mancino V, Slepak T, Tachiiri Y, Simon M (1992) Proc Natl Acad Sci USA 89:8794–8797 55. Schleper C, DeLong EF, Preston CM, Feldman RA, Wu KY, Swanson RV (1998) J Bacteriol 180:5003–5009 56. Beja O, Spudich EN, Spudich JL, Leclerc M, DeLong EF (2001) Nature 416:786–789 57. Beja O, Koonin EV, Aravind L, Taylor LT, Seitz H, Stein JL, Bensen DC, Feldman RA, Swanson RV, DeLong EF (2002) Appl Environ Microbiol 68:335–345 58. Gill I, Valivety R (1997) TIBTECH 15:401–409 59. Yazawa, K (1996) Lipids 31:297–300 60. Takeyama H, Takeda D, Yazawa K, Yamada A, Matsunaga T (1997) Microbiology 143:2725–2731 61. Nystrom T, Flardh K, Kjelleberg S (1990) J Bacteriol 172:7085–7097 62. Nystrom T, Olsson RM, Kjelleberg S (1992) Appl Environ Microbiol 58:55–65 63. Gross M, Kosmowsky IJ, Lorenz R, Molitoris HP, Jaenicke R (1994) Electrophoresis 15:1559–1565 64. Rabus R, Gade D, Helbig R, Bauer M, Glockner FO, Kube M, Schlesner H, Reinhardt R, Amann R (2002) Proteomics 2:649–655 65. Doino Lemus J, McFall-Ngai MJ (2000) Appl Environ Microbiol 66:4091–4097 66. Lopez JL, Marina A, Alvarez G, Vazquez J (2002) Proteomics 2:1658–1665 67. Okamoto OK, Hastings JW (2003) Gene 321:73–81 68. Peplies J, Glockner FO, Amann R (2003) Appl Environ Microbiol 69:1397–1407 69. Liu W-T, Mirzabekov AD, Stahl DA (2001) Environ Microbiol 3:619–629

46

T. Schweder et al.

70. Wu L, Thompson DK, Li G, Hurt RA, Tiedje JM, Zhou J (2001) Appl Environ Microbiol 67:5780–5790 71. Rosamond J, Allsop A (2000) Science 287:1973–1976 72. Jones RN, Pfaller MA (1998) Diagn Microbiol Infect Dis 31:379–388 73. File TM Jr (1999) Chest 115:3–8 74. Dobrindt U, Blum-Oehler G, Nagy G, Schneider G, Johann A, Gottschalk G, Hacker J (2002) Infect Immun 70:6365–6372 75. Oelschlaeger TA, Dobrindt U, Hacker J (2002) Int J Antimicrob Agents 19:517–521 76. Hacker J, Hentschel U, Dobrindt U (2003) Science 301:790–793 77. Chen CY, Wu KM, Chang YC, Chang CH, Tsai HC, Liao TL, Liu YM, Chen HJ, Shen AB, Li JC, Su TL, Shao CP, Lee CT, Hor LI, Tsai SF (2003) Genome Res 13:2577– 2587 78. Gerth K, Pradella S, Perlova O, Beyer S, Müller R (2003) J Biotechnol 106:233–253 79. Jensen PR, Fenical W (1994) Annu Rev Microbiol 48:559–584 80. Bernan VS, Greenstein M, Maiese WM (1997) Adv Appl Microbiol 43:57–90 81. Rappe MS, Connon SA, Vergin KL, Giovannoni SJ (2002) Nature 418:630–633 82. Button DK, Schut F, Quang P, Martin R, Robertson BR (1993) Appl Envir Microbiol 59:881–891 83. Eder W, Jahnke LL, Schmidt M, Huber R (2001) Appl Envir Microbiol 67:3077–3085 84. Lee N, Nielsen PH, Andreasen KH, Juretschko S, Nielsen JL, Schleifer KH, Wagner M (1999) Appl Environ Microbiol 65:1289–1297 85. Bruns A, Cypionka H, Overmann J (2002) Appl Environ Microbiol 68:3978–3987 86. Burja AM, Abou-Mansour E, Banaigs B, Payri C, Burgess JG, Wright PC. J (2002) Microbiol Methods 48:207–219 87. Slattery M, Rajbhandari I, Wesson K (2001) Microb Ecol 41:90–96 88. Faulkner DJ (2000) Antonie van Leeuwenhoek 77:135–145 89. Pfefferle CM, Suessmuth RD (2003) Screening 5:41–43 90. Schmid I, Sattler I, Grabley S, Thiericke R (1999) BIOforum 1–2:17–20, and 5:261– 264 91. Grabley S, Thiericke R, Zeeck A (1999) In: Grabley S, Thiericke R (eds) Drug discovery from nature. Springer, Berlin Heidelberg New York, pp 124–148 92. Clerval R, Kühner M, Li Z, Minas W, Fjällman A, Witholt B, Duetz WA (2000) BioWorld 6:24–26 93. Hamburger M, Danz H, Dittmann K (2002) Screening 2:33–35 94. Bhakuni DS (1998) J Indian Chem Soc 75:191–205 95. Grabley S, Thiericke R (1999) In: Grabley S, Thiericke R (eds) Drug discovery from nature. Springer, Berlin Heidelberg New York, pp 38–48 96. Frobel K, Metzger S (2001) In: Rehm HJ, Reed G (eds) Biotechnology, vol 10: special processes. Wiley VCH, pp 41–60 97. Vogel HG (ed) (2000) Drug discovery and evaluation, 2cnd edn. Springer, Berlin Heidelberg New York 98. Minor L (2003) Screening 5:25–28 99. Jensen PR, Fenical W (1994) Ann Rev Microbiol 48:559–584 100. Devlin JP (2002) In: Mei HY, Czarnik AW (eds) Integrated drug discovery technologies. Marcel Dekker, New York, pp 221–245 101. Lebl M (2002) In: Mei HY, Czarnik AW (eds) Integrated drug discovery technologies. Marcel Dekker, New York, pp 395–405 102. Otrocka A, Oehlenschlager F, de Hoop M (2003) Screening 1:41–43 103. Bertels S, Frormann S, Jas G, Bindseil K (1999) In: Grabley S, Thiericke R (eds) Drug discovery from nature. Springer, Berlin Heidelberg New York, pp 72–105

Screening for New Metabolites from Marine Microorganisms

47

104. Okun I, Kaler G et al. (2000) SPIE Proc 3921:90–100 105. Edwards BS, Kuckuck FW, Prossnitz ER, Ransom JT, Sklar LA (2001) J Biomol Screen 6:83–90 106. Shapiro HM (1995) Practical flow cytometry. Wiley-Liss, New York 107. Lindon JC, Bailey NJC, Nicholson JK, Wilson, ID (2003) Handbook of analytical separations, vol 4: bioanalytical separations. pp 293–329 108. Lindon JC, Nicholson JK, Wilson ID (2000) J Chromatogr B 748:233–258 109. Fenical W (1993) Chem Rev 93:1673–1683 110. Jensen PR, Fenical W (2000): In: Fusetani N (ed) Drugs from the Sea. Karger, Basel, pp 6–29 111. Eder C, Proksch P (1997) DROGENREPORT 10:24–28 112. Feling RH, Buchanan GO, Mincer TJ, Kauffman CA, Jensen PR, Fenical W (2003) Angew Chem 115:369–371 113. Shigemori H, Bae MA, Yazawa K, Sasaki T, Kobayashi J (1992) J Org Chem 57:4317– 4320 114. Varoglu M, Corbett TH, Valeriote FA, Crews P (1997) J Org Chem 62:7078–7079 115. Takahashi C, Takada T, Yamada T, Minoura K, Uchida K, Matsumura E, Numata A (1994) Tetrahedron Lett 35:5013–5014 116. Baz JP, Canedo LM, Fernan’dez Puentes JL, Silva Elipe MV (1997) J Antibiot 50:738– 741 117. Takahashi C, Numata A, Ito Y, Matsumura E, Minoura K, Eto H, Shingu T, Ito T, Hasegawa T (1994) J Antibiot 47:1242–1249 118. Renner MK, Jensen PR, Fenical W (1998) J Org Chem 63:8346–8354 119. Trimurtulu G, Ohtani I, Patterson GML, Moore RE, Corbett TH, Valeriote FA, Demchik L (1994) J Am Chem Soc 116:4729–4737 120. Gerwick WH, Proteau PJ, Nagle DG, Hamel E, Blokhin A, Slate DL (1994) J Org Chem 59:1243–1245 121. Luesch H, Yoshida WY, Moore RE, Paul VJ, Mooberry SL, Corbett T (2002) J Nat Prod 65:16–20 122. Luesch H, Yoshida WY, Moore RE, Paul VJ (1999) J Nat Prod 62:1702–1706 123. Gerard J, Lloyd R, Barsby T, Haden P, Kelly MT, Andersen RJ (1997) J Nat Prod 60:223–229 124. Pathirana C, Jensen PR, Fenical W (1992) Tetrahedron Lett 33:7663–7666 125. Höller U, König GM, Wright AD (1999) J Nat Prod 62:114–118 126. Kusnick C, Jansen R, Liberra K, Lindequist U (2002) Pharmazie 57:510–512 127. Liberra K, Jansen R, Lindequist U (1998) Pharmazie 53:578–581 128. Schlingmann G, Milne L, Williams DR, Carter GT (1998) J Antibiot 51:303–316 129. Gustafson K, Roman M, Fenical W (1989) J Am Chem Soc 111:7519–7524 130. Davidson BS, Schumacher RW (1993) Tetrahedron 49:6569–6574 131. Matsuda M, Shigeta S, Okutani K (1999) Mar Biotechnol 1:68–73 132. Trischman A, Tapiolas DM, Jensen PR, Dwight R, Fenical W (1994) J Am Chem Soc 116:757–758 133. Kawano Y, Asada M, Inoue M, Nakagomi K, Oka S, Higashirana T (1998) J Mar Biotechnol 6:49–52 134. Sugano M, Sato A, Iijima Y, Oshima T, Furuya K, Kuwano H, Hata T, Hanzawa H (1991) J Amer Chem Soc 113:5463–5464 135. Takaishi S, Tuchiya N, Sato A, Negishi T, Takamatsu Y, Matsushita Y, Watanabe T, Iijima Y, Haruyama H, Kinoshita T, Tanaka M, Kodama K (1998) J Antibiot 51:805– 815 136. Poch GK, Gloer JB (1989) Tetrahedron Lett 30:3483–3486

48

T. Schweder et al.

137. Itoh T, Kinoshita M, Aoki S, Kobayashi M (2003) J Nat Prod 66:1373–1377 138. Kakeya H, Takahashi I, Okada G, Isono K, Osada H (1995) J Antibiot 48:733–735 139. Han B, McPhail KL, Ligresti A, Di Marzo V, Gerwick WH (2003) J Nat Prod 66:1364– 1368 140. Bandow JE, Brötz H, Leichert LOI, Labischinski H, Hecker M (2003) Antimicrob Agents Chemother 47:948–955 141. Sender U, Bandow EJ, Engelmann S, Lindequist U, Hecker M (2004) Pharmazie 59:65–70 142. Lindequist U, Sender U, Bandow J, Jülich WD, Kusnick C, Schweder T, Hecker M (2001) Antibacterial substances from marine fungi and their investigation by proteome-based methods. In: Proceedings of 8th international marine and freshwater mycology symposium. Hurghada, Egypt, 2001, p 457 143. Bode BH, Bethe B, Höfs R, Zeeck A (2002) Chem Bio Chem 3:619–627

Adv Biochem Engin/Biotechnol (2005) 96: 49–125 DOI 10.1007/b135782 © Springer-Verlag Berlin Heidelberg 2005 Published online: 24 August 2005

Fatty Acids from Lipids of Marine Organisms: Molecular Biodiversity, Roles as Biomarkers, Biologically Active Compounds, and Economical Aspects Jean-Pascal Bergé1 (u) · Gilles Barnathan2 1 Centre

de Nantes, Laboratoire Génie Alimentaire, Département Valorisation des Produits, Institut Français pour l’Exploitation de la Mer (IFREMER), BP21105, 44311 Nantes Cedex 03, France [email protected] 2 Pˆ ole Mer et Littoral, Laboratoire de Chimie Marine, Groupe SMAB (EA 2160), Substances marines à activité biologique, Université de Nantes, 2 rue La Houssinière, 44322 Nantes Cedex 03, France [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Marine bacteria and cyanobacteria . . . . . . . . . . . . . . . . . . . . .

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Marine invertebrates . . Sponges . . . . . . . . . ∆5,9 fatty acids . . . . . Branched fatty acids . . Methoxylated fatty acids Acetylenic fatty acids . . Coelenterate – Cnidaria . Echinodermata . . . . . Tunicates . . . . . . . . . Molluscs . . . . . . . . . Introduction . . . . . . . Mussels . . . . . . . . . . Oysters . . . . . . . . . . Patella . . . . . . . . . . Clams . . . . . . . . . . Scallops . . . . . . . . . Squids . . . . . . . . . .

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78 78 79 80 80 81 81 81 81 82 82 82 83 86 87 90 90 103 105 105 111

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10 10.1 10.2 10.2.1 10.2.2 10.2.3 10.2.4 10.2.5 10.2.6 10.2.7 10.2.8 10.3 10.4 10.4.1 10.4.2 10.5 10.5.1 10.5.2 10.6 10.6.1 10.6.2

Polyunsaturated FA (n-3) of commercial interest . . . . . . . . . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Health benefits . . . . . . . . . . . . . . . . . . . . . . . . . . . . Heart health . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cancer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Arthritis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Psoriasis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lung disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Attention-deficit disorder . . . . . . . . . . . . . . . . . . . . . . . Mental health . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pregnancy and infancy . . . . . . . . . . . . . . . . . . . . . . . . Nutrition: importance of the ratio of (n-6) and (n-3) essential FA Routes for biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . Aerobic pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . Anaerobic pathway . . . . . . . . . . . . . . . . . . . . . . . . . . Some promising sources of marine LC-PUFA . . . . . . . . . . . . PUFA from nonphotosynthetic microorganisms . . . . . . . . . . PUFA from fish . . . . . . . . . . . . . . . . . . . . . . . . . . . . Current utilization of marine oils and lipids . . . . . . . . . . . . Market . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Common resources . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract Because of their characteristic living environments, marine organisms produce a variety of lipids. Fatty acids constitute the essential part of triglycerides and wax esters, which are the major components of fats and oils. Nevertheless, phospholipids and glycolipids have considerable importance and will be taken into account, especially the latter compounds that excite increasing interest regarding their promising biological activities. Thus, in addition to the major polyunsaturated fatty acids (PUFA) such as eicosapentaenoic (EPA) and docosahexaenoic (DHA) acids, a great number of various fatty acids occur in marine organisms, e.g. saturated, mono- and diunsaturated, branched, halogenated, hydroxylated, methoxylated, non-methylene-interrupted. Various unprecedented chemical structures of fatty acids, and lipid-containing fatty acids, have recently been discovered, especially from the most primitive animals such as sponges and gorgonians. This review of marine lipidology deals with recent advances in the field of fatty acids since the end of the 1990s. Different approaches will be followed, mainly developing biomarkers of trophic chains in marine ecosystems and of chemotaxonomic interest, reporting new structures, especially those with biological activities or biosynthetic interest. An important part of this review will be devoted to the major PUFA, their relevance to health and nutrition, their biosynthesis, their sources (usual and promising) and market. Keywords Lipids · Fatty acids · Marine organisms · Biomarkers · Nutrition Abbreviations AA arachidonic acid 20:4(n-6) DHA docosahexaenoic acid 22:6(n-3) EPA eicosapentaenoic acid 20:5(n-3) FA fatty acid

Fatty acids from marine I FATM GC/MS PUFA LC-PUFA VLC-PUFA MUFA NMI NMID PL SFA TL EFA TAG SCO

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fatty-acid trophic markers gas-chromatography mass spectrometry polyunsaturated fatty acids long-chain polyunsaturated fatty acids Very-long-chain polyunsaturated fatty acids monounsaturated fatty acids non-methylene-interrupted fatty acids non-methylene-interrupted dienoic fatty acids phospholipids saturated fatty acids total lipids essential fatty acid triacylglycerols single-cell oil

1 Introduction Lipids are major sources of metabolic energy and essential materials for the formation of cell and tissue membranes. They are very important in the physiology and reproductive processes of marine animals and reflect the special biochemical and ecological conditions of the marine environment [1–3]. The interest of chemists, biochemists and biotechnologists in lipids and fatty acids (FA) from marine animals and algae has been stimulated, in particular, by the recognition that polyunsaturated fatty acids (PUFA) are important to human health and nutrition. They are required for reproduction and growth. The relative proportion and composition of FA in marine organisms are characteristic for every species and genus, and also depend on environmental conditions. Several comprehensive reviews are available on marine FA, their occurrence, their roles and the methods used in their analysis [4–8]. The principal role of neutral lipids, which in marine organisms consist predominantly of triacylglycerols (TAG) and wax esters, is as an energetic reserve of FA that are destined either for oxidation to provide energy (ATP) or for incorporation into phospholipids. Phospholipids are the building blocks for the membrane lipid bilayer. FA provide the hydrophobic interior of all cell membranes, forming an impermeable barrier to water and polar molecules and separating the cell contents from the extracellular medium. The physical properties of the membrane are determined by the individual lipids within the FA components of the lipids and their interaction with sterols and proteins. While their function as structural lipids in membranes has been known for a long time, ceramides and glycosyl ceramides (glycosphingolipids) play an important role in many fields of cell biochemistry, such as molecular recognition. In addition, ceramides from marine organisms have excited great attention as signal transducers, and some of them have

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been recognized as possessing antimicrobial and cytotoxic activities. Marine glycosphingolipids, chiefly isolated from sponges, show interesting biological activities such as immunomodulation and antitumoral activity [9]. The fatty acyl chains linked to these classes of compounds are often commonly occurring but several new and original structures have been reported recently [10, 11]. Arising from alpha-glycosphingolipids isolated from marine sponges, the simplest (KRN 7000) is being considered as an anticancer agent ([12], see Bourguet-Kondracki & Kornprobst, this book). Fat not only provides energy, it facilitates the absorption of fat-soluble vitamins (vitamins A, D, E, and K), and plays an important role in the production and regulation of eicosanoids. In addition, lipid class and FA compositions can be used to understand and identify food web interactions. Different approaches can be applied in this field of marine lipidology, focused on FA, mainly: – – – – –

searching for new FA structures evaluating new sources of major PUFA of biological interest evaluating their role in cell membranes investigating biosynthetic pathways developing trophic and/or chemotaxonomic biomarkers in ecosystems.

The rapid development of excellent analytical methods, especially gas liquid chromatography coupled to mass spectrometry (GC/MS), which can deal with complex mixtures, has also been a major contributing factor to the progress in the chemistry and biochemistry of marine lipids [5, 6, 13]. As the principal producers in the marine environment, microalgae support both pelagic and benthic food webs, and their lipids and FA are been extensively studied. Microalgae are known to have different FA compositions depending on their taxonomic position [14, 15]. At the next trophic level, zooplankton form an essential link between primary producers and higher-order consumers. Very recently, a complete review was devoted to fatty-acid trophic markers (FATM) in the pelagic marine environment [8]. The FATM concept is based on the observation that marine primary producers lay down certain FA patterns that may be conservatively transferred through aquatic food webs. Thus, they can be recognized in their primary consumers. The next step is concerned with the dynamics of FA in fish, which catabolize and transform dietary FA. Marine invertebrates, especially sponges, have proved to be a rich source of many unusual FA. Sponges are very ancient animals with special structural features, particularly the cell membrane, which have allowed their adaptation to often precarious environments. These primitive organisms are difficult to classify due to the few available useful morphological characteristics. The use of taxonomic knowledge allows investigation to be focused on genera that offer the great potential in metabolites of biological interest. Recently, a comprehensive taxonomy was published, which provides the state of the art [16]. Representatives of the Coelenterate phylum have remarkable peculiarities in

Fatty acids from marine I

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their FA composition. Some species contain unusual FA or unusual concentrations of common components. The main purposes of this paper are to illustrate the molecular biodiversity in marine lipids (from the first planktonic marine producers to fish) and, in another part, to focus on the most important PUFA (sources, biosynthesis, economy). This work has been done by considering the most interesting advances in marine lipidology since the end of the 20th century. Thus, without covering the subject exhaustively, this review deals with FA from total lipids or those specifically linked to a particular lipid class such as TAG, wax esters, ceramides, polar lipids and some additional atypical secondary metabolites. Thus, unusual or novel FA, and FA that are interesting due to their applications in chemotaxonomy, biosynthesis and as biomarkers, will mainly be taken into account.

2 Nomenclature of fatty acids According to the international nomenclature, the position of the first double bond is given by the (n-x) notation, counting the number of carbon atoms from the methyl end. For instance, 18:4(n-3) identifies an FA with 18 carbon atoms and four double bonds, the first double bond occurring after the third carbon atom [4, 7] (Fig. 1). An additional PUFA is depicted in Fig. 1 (middle), namely 24:6(n-3). According to an alternative notation, the locations of the double bonds are counted from C-1 (the carboxyl group). Thus, 18:4(n-3) is designated 6,9,12,15-18:4. This latter notation will be used in particular in the case of non-methylene-interrupted fatty acids (NMI FA), such as 7,13,17-20:3 shown above, the most commonly encountered among these being the dienoic acids (NMID) (Fig. 1, bottom). The configuration of double bonds, generally assumed as cis (Z) in natural compounds, must be indicated in other cases. The positions of a methyl branch or another group is indicated by the number of the carbon atom on which the chain is substituted (e.g. 10-methylhexadecanoic acid or simply 10-Me-16:0; 2-hydroxydocosanoic acid or 2-OH-22:0 ; 6-bromo-5,9heptacosadienoic acid or 6-bromo-5,9-27:2).

3 Fatty-acid biosynthetic pathways in primary producers The basic processes of FA biosynthesis are summarized in Fig. 2. The de novo biosynthesis of FA follows the common pathway with major end products

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Fig. 1 Examples of numbering and designation of fatty acids (FA)

being mainly 16:0 and also 14:0, 18:0 and 20:0 (also produced by chain elongation) [8]. Then, an aerobic desaturation is catalyzed by the enzyme ∆9-desaturase to give rise to 16:1(n-7), 18:1(n-9) and 20:1(n-11). Generally, only plants are capable of biosynthesizing de novo (n-3) and (n-6) PUFA (Fig. 2A). Oleic acid 18:1(n-9) is the precursor of all (n-3) and (n-6) PUFA. The next double bonds are introduced to form 18:2(n-6) and 18:3(n-3). Through appropriate desaturations and chain elongations, 18:2(n-6) may be further converted to 20:4(n-6) (AA), and 18:3(n-3) to EPA. DHA is obtained via C24 PUFA intermediates rather than direct elongation of EPA, according to the so-called Sprecher pathway [21–23]. This biosynthetic scheme is typically observed in dinoflagellates, in which FA such as 18:4(n-3) and DHA are often dominant. Furthermore, the biosynthetic pathway producing 16:4(n-1) from 16:0 is characteristic of diatoms [8, 24]. The de novo biosynthesis of long-chain monounsaturated FA (MUFA) typically pronounced in calanoid copepods is showed in Fig. 2B. Biosynthetic considerations will be developed in detail in the last part of this review.

4 Marine bacteria and cyanobacteria Marine bacteria are known for their role in nutrient cycling and the degradation of organic matter [8]. The roles of bacteria in marine food webs has two aspects, firstly as primary food sources, and secondly as components of the commensal microbial communities of marine animals [25]. Marine heterotrophic bacteria are abundant in sediments and as colonizers of settling particulate matter following plankton blooms [26]. That observation explains why the FA composition of marine bacteria has mainly been studied by geochemists [26–29]. Bacteria incorporate FA mainly in PL. Bacterial FA are

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Fig. 2 Major pathways of FA biosynthesis in marine algae (a), modified after Gurr & Harwood [17] and Cook [18], and herbivorous calanoid copepods (b), modified after Sargent & Henderson [19] and Kattner & Hagen [20]. Extracted from Dalsgaard et al. [8]

commonly saturated (SFA) and monounsaturated (MUFA), ranging from C10 to C20 , whereas PUFA are quite rare. Bacterial FA biomarkers are typically odd-numbered, branched trans-unsaturated and cyclopropyl FA such as 15:0, 17:0, iso- and anteiso-branched SFA and MUFA, 10-methyl-16:0 ([8] and references therein). Furthermore, deep-sea bacteria and several bacterial strains (Pseudomonas, Vibrio) have been shown to be capable of producing (n-3) PUFA, as recently reviewed [24, 25, 30–32]. Very little is known about the biosynthesis of PUFA in bacteria. EPA and DHA in bacteria are contained within phospholipids rather than TAG. Therefore, marine bacteria seem to be of limited use as a source of oils rich in (n-3) PUFA [24]. (see the chapter entitled “Promising sources of marine LC-PUFA”) Cyanobacteria, a class of photosynthetic prokaryotes occurring in the phytoplankton, produce C18 PUFA esterified to polar lipids, but they do not biosynthesize EPA or DHA [24]. EPA production was obtained by a transgenic marine cyanobacterium carrying a plasmid containing the essential open reading frames for EPA synthesis [33].

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Fig. 3 Malingamide G and its potential precursor 7-methoxy-dodec-4-enoic acid

Fatty-acid amides are widespread in nature [34]. They are incorporated into some lipid classes such as ceramides, glycosphingolipids and various Nacylated lipid molecules. A new diacylgalactolipid was isolated from the marine cyanobacterium Oscillatoria sp. comprised of 9,12-octadecadienoyl and 4-hexadecenoyl chains [35]. Cyanobacteria of the genus Lyngbya are a rich source of bioactive secondary metabolites including fatty-acid amides [36]. A series of biologically active malyngamides has been identified from marine cyanobacteria with a mid-chain methoxylated fatty acyl chain [37–40]. As an example, Malyngamide G and the new 7-methoxydodec-4(E)-enoic acid (its possible precursor) were isolated from Lyngbya majuscula collected off the French Mediterranean coast (Fig. 3). Both these compounds are non-cytotoxic to KB cells and show immunosuppressive activity [39]. In several other malyngamides, the fatty-acid amid is 7-methoxy-tetradecen-4-enoic.

5 Phytoplankton Primary producers provide the basic FA patterns in marine food webs. They consist of macroalgae and phytoplankton, which mainly comprise microalgae and photoautotrophic bacteria. Phytoplankton in the pelagic environment comprises mainly Bacillarophyceae (diatoms), Dinophyceae (dinoflagellates) and Prymnesiophyceae. Algal FA are biosynthesized in the chloroplasts comprising the thylakoid membranes, and are chiefly esterified to glycolipids rich in (n-3) PUFA. During the exponential growth phase of phytoplankton blooms, carbon fixed through photosynthesis is allocated to growth and cell division rather than lipid storage. Therefore, the level of glycolipids is particularly high in this phase, and the proportion of (n-3) PUFA can reach 50% of total lipids [8, 14, 41, 42]. It is well known that plants are usually the only organisms that can biosynthesize de novo the acids 18:2(n-6) and 18:3(n-3). These FA and their principal derivatives (e.g. AA, EPA and DHA) are essential constituents of heterotrophic organisms. Thus, algae occupy a central position within marine food webs. As shown in a recent review, FA patterns can be used as potential taxonomic markers regarding the presence and combinations of certain FA that

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can be characteristic of particular algal classes, whereas individual FA cannot be used as indicators of particular algal species [43]. This approach has been developed in the case of diatoms and dinoflagellates, two important classes in marine environments. Thus, high values of 16:1(n-7)/16:0 (typically

> 1), and C16 / C18 have been associated with a dominance of bacillariophytes [28, 29, 44]. Furthermore, high values of 18:5(n-3)/18:3(n-3) and
(C18 PUFA, C22 PUFA) have been associated with dinophytes [45]. Bacillariophytes

can be distinguished from dinophytes by means of high values of C16 / C18 together with low values of 18:5(n-3)/18:3(n-3) [45]. This criteria can be reinforced by the examination of the ratio 22:6(n-3)/20:5(n-3) [28]. A value ≥ 1 indicates a predominance of dinophytes, while a value < 1 indicates a predominance of bacillariophytes. The lipids of diatoms, characterized by high levels in EPA and the absence of DHA, are also rich in C16 PUFA [24]. If investigations of FA isolation and purification have principally been carried out on (n-3) PUFA, there are other potentially interesting FA commercially unavailable. Thus, the acids 16:3(n-4), 16:2(n-4) and 16:2(n-7) were isolated as methyl esters by means of liquid chromatography using a porous graphitic-carbon phase [46]. The taxonomy of the Raphidophyceae is still uncertain and needs the help of chemotaxonomic data, such as FA composition, in order to distinguish correctly the genera [45–49]. The FA compositions of twelve raphidophyte strains were established and were very similar to previous data [47]. The major PUFA were EPA (14.8–24.5%) and 18:4(n-3) (12.0–26.6%, with an exception at 0.3%) [49]. High levels of free FA were observed in lipids of Fibrocapsa species (23.6–37.9%). FA profiles allowed clear discrimination between the genera. By using a selected EPA-deficient mutant of Porphyridium cruentum, it has been demonstrated that TAG of the red microalga P. cruentum can contribute to the biosynthesis of eukaryotic galactolipids [50]. FA biomarkers were used to investigate the biogeochemistry of a former blue-mussel aquaculture site and the high levels of PUFA found indicated a substantial phytoplankton source (diatoms and dinoflagellates) [29]. FA composition of toxic microalgae have been determined to detect useful biomarkers in screening seawater seafood samples [51]. Two Pseudo-nitzschia species were studied and both displayed similar FA compositions typical of diatoms, including 16:1(n-7), 16:2(n-4) and EPA as major unsaturated FA. 16:4(n-1) occurred in both species and therefore could be used as a signature compound in differentiating toxic Pseudo-nitzschia from other diatoms [51]. Furthermore, the possibility of using boiling water to deactivate lipolytic enzymes, as previously found [52], was confirmed, and it was suggested that some mechanisms of PUFA degradation was also inhibited [51]. The FA compositions of microalgae have been shown to change in response to changes in salinity. Green unicellular microalgae of the genus Dunaliella are known for their capability to grow at high salinities up to salt-saturated water [53]. The major unsaturated FA in two Dunaliella iso-

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lates originating from Antarctic hypersaline lakes were 16:4(n-3), 18:3(n-3), 18:2(n-6), 18:1(n-9) and 16:1(n-7). The results suggest that the appropriate environment necessary for the growth of these halophilic species is a certain level of osmotic pressure in the medium [53]. High levels of DHA are noticeable in the lipids of the Dinophyceae, another major component of marine phytoplankton, together with significant amounts of EPA, 18:5(n-3) and 18:4(n-3) [24, 54]. Thus, it seems possible that dinoflagellates are capable of synthesizing DHA through the chainshortening of 24:6(n-3), since a similar mechanism could produce 18:5(n-3) from EPA [24]. The presence of EPA and also DHA and 18:5(n-3) has been linked to potential toxicity in a raphidophyte Heterosigma as well as in a dinoflagellate Gymnodinium [55, 56]. It was suggested that the mode of action in the ichthyotoxicity of these harmful bloom-forming flagellates is correlated to oxygen radical formation [49]. Two Gymnodinium species and several other dinoflagellate species were found to contain unusual octacosapolyenoic FA, namely 28:7(n-6) and 28:8(n-3) at levels up to 2.2% of total FA [57] (Fig. 4). VLC-PUFA have also been found in some autotrophic and heterotrophic lower organisms such as microalgae, fungi, sponges and bacteria [59], and most of them are either saturated or monounsaturated. Such FA were identified in Baltic herring, namely 28:7(n-3) and 28:7(n-6) [58]. Furthermore, the two Prorocentrum species studied were found to contain 18:4(n-3) (12.7–15.3%), 18:5(n-3) (36.4–37.6%) and DHA (18.3–22.0%) [57, 58]. 28:8 (n-3) was also identified in a commercially available oil, used as dietary supplements of PUFA for humans, from the heterotrophic dinoflagellate Cryptecodinium cohnii [60]. Furthermore, these unusual octacosapolyenoic FA also occurred as minor components in lipids of all five dinoflagellates studied for fatty acid and sterol compositions [57]. The major unsaturated FA were as followed: 18:4(n-3) (2.5–11.5%), 18:5(n-3) (7.0–43.1%), EPA (1.8–20.9%) and DHA (9.9–26.3%). A new ceramide isolated from the epiphytic dinoflagellate Coolia monotis bears a novel fatty acyl moiety, namely 2-hydroxy-15-methyl3-octadecenoyl [61] (Fig. 5). The fatty acid composition of polar lipids and TAG was determined in different types of symbiotic dinoflagellates isolated from several hermatypic corals from a fringing reef in Japan [15]. 18:4(n-3) (10.0–26.2%) and DHA

Fig. 4 Octacosapolyenoic FA from dinoflagellates

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Fig. 5 New ceramide from the dinoflagellate Coolia monolis

(10.6–17.8%) were found as major PUFA in polar lipids of symbiotic dinoflagellates isolated from all species studied. In addition, polar lipids of dinoflagellates from Millepora intricata were different from those originating from other corals in that they contained high amounts of 18:5(n-3) (8.7%) and 22:5(n-6) (10.3%). It was suggested that FA might provide useful information on possible taxonomic differences among symbiotic dinoflagellates [15].

6 Macroalgae FA in marine algae have attracted considerable attention among researchers because they can produce significant amounts of interesting PUFA and provide useful distinguished features of chemotaxonomic value [62]. The data available on lipids from macroalgae have been reviewed recently [8]. Three classes are mainly concerned: Chlorophyceae, Rhodophyceae and Phaeophyceae. FA from 11 macroalgae from the French Brittany coast were studied [63]. More recently, Li et al. [62] reported the FA compositions of 22 species of marine macrophytes, collected from the coast of the Bohai Sea belonging to the three aforementioned algal classes. These algae have FA patterns typical of red, brown and green algae from other regions. In general, red algae from the Bohai Sea contained high levels of C20 PUFA, primarily that of EPA (up to 37.5%) and AA (up to 29.4%). The main difference in the FA compositions between red and brown algae was that the latter were richer in C18 PUFA, especially in 18:4(n-3) (up to 20.1%). Seven of the teen brown algal species studied also contained EPA as a major component, accounting for 8.4–24.2%. Green algae studied had the highest level of C18 PUFA, mainly 18:3(n-3) (20.5–27.2% of total lipids) and 18:4(n-3), and the lowest level of C20 PUFA [62]. Interestingly, another study of FA composition of 19 species of the same algal classes, collected on the Pacific coast of North California gave rise to very similar conclusions [64]. Red Californian algae contained AA (5.3–23.4%) and EPA (27.8–45.4%). Brown algae contained 18:4(n-3) (3.6–18.6%) and EPA (3.1–15.5%). Two of the three green algae studied contained 16:4(n-3) (13.6–16.2%) and 18:4(n-3) (12.1–22.1%). Both these studies show that red, brown and green algae have distinguished FA

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Fig. 6 New methoxylated FA from lipids of Schyzimenia dubyi

profiles that do not depend on the geographical location of the algae and that have a chemotaxonomic significance for seaweeds [62, 64]. A recent comparative study of FA composition of Arctic and Antarctic macroalgae considered their use as indicators of phylogenetic and trophic relationships [65]. Several eicosanoids, metabolites of AA, such as hydroxytetraenoic acids, associated with prostaglandins, were identified in a Japanese red alga Gracilaria asiatica [66]. Methoxy FA are not very widespread in nature [67, 68]. Several mid-chain methoxy FA have been reported only in certain microorganisms and marine cyanobacteria (genus Lyngbya), as seen above. Four novel mid-chain methoxy FA (16% of total acid mixture) were identified in lipids from a red alga as 9-MeO-15:0, 9-MeO-17:0, 13-MeO-21:0 and 15-MeO-23:0 acids [69], as depicted in Fig. 6. These algal lipids contained C14 -C28 SFA, accounting for 77%, and hydroxylated FA, but no PUFA. Furthermore, marine sponges have provided new 2-methoxy long-chain acids that will be presented in a following chapter.

7 Zooplankton During the last decade, zooplanktonic organisms from Arctic and Antarctic waters have given rise to intense research, especially on lipids [70–90]. These investigations have highlighted the general lipid characteristics in high-latitude zooplankton communities, such as copepods and ctenophores, in terms of food webs and biomarkers. The Southern Ocean has a complex food web including planktivorous herbivores (krill, salps, copepods) that are fed upon by fish, squid, seals and whales [83, 87]. Krill (Euphausia superba) provide 30–90% of the diet for these carnivores and have an estimated

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standing stock biomass of 200–400 million metric tons. This high biomass reflects krill’s ability to adapt to marked seasonality in food supply. Krill are primarily herbivores feeding on phytoplankton in the summer. In the winter krill feed mainly on ice algae [87, 89]. Krill lipids have been intensively studied because of their commercial interest and undisputed importance in the Southern Ocean, while the importance of gelatinous organisms, such as salps, ctenophores and medusae, is now recognized in marine pelagic ecosystems. As the best-studied group of zooplankton with respect to the FA trophicmarkers concept, herbivorous calanoid copepods have been mainly examined. They dominate the zooplankton biomass in high-latitude ecosystems and accumulate large lipid reserves. In addition, the FA characteristics of omnivorous and carnivorous copepods have been summarized, especially using FA as markers of carnivory [8]. Moreover, calanoid copepods are themselves important producers of specific FA and fatty alcohols (from wax esters). They play an important role in polar food webs and provide higher trophic levels with a lipid-rich high-energy diet [71]. However, similarities and differences between species emerge more clearly if the major lipid classes are analysed separately to determine their compositions. Phospholipids and the storage lipids (TAG and wax esters) are expected to exhibit strong compositional differences, which may provide additional information on their structural and energetic functions [71]. The compositions of wax esters, TAG and phospholipids in nine Arctic and Antarctic copepods have provided evidence of energetic adaptations with similarities and differences between the species [71]. The wax esters of the herbivorous species were clearly characterized by the long-chain monounsaturated FA 20:1(n-9) and 22:1(n-11), whereas the omnivorous and carnivorous species usually had high relative amounts of 18:1(n-9). The phospholipids contained very high levels of PUFA, especially 22:6(n-3). The phospholipid FA compositions in both Arctic and Antarctic species were found to be very similar. This extremely high degree of unsaturation (EPA and DHA together accounted for 46–60% of the total phospholipid FA) is quite unusual. In a recent investigation, the variation in the FA content was related to the spatial distribution of krill and the available diet as well as to maturity and sex [83]. E. superba are known as essentially herbivores when phytoplankton is abundant, but they can be omnivorous if algal biomass becomes relatively low. Three regionally groups of krills were considered. Krills from one group, almost exclusively juveniles, were surviving in the region characterized by lowest algal biomass and had probably resorted to carnivory on PUFA-rich copepods [83]. The latter krill had unusually high level of PUFAs, mainly 18:4(n-3), EPA and DHA. Changes in lipid composition of the Antarctic E. superba were investigated regarding the influence of geographical location, the sexual maturity stage and the distribution among organs [73].

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Lipid metabolism of E. crystallorophias and its ecological implications have been studied because this euphausiid is the dominant krill species in highAntarctic waters. Thus, it has to cope with the most extreme environmental conditions of all euphausiids [76]. Wax esters were the primary storage lipids and accounted for up to 55.6% of total lipids, including 18:1(n-9) and 18:1(n-7) as major components (together up to 90% of total wax esters acids). 16:0, EPA, DHA and 18:1(n-9) were the major phospholipid FA components. Little is known about lipids and FA of a member of gelatinous zooplankton, the salps (Tunicates). The pelagic tunicate Dolioletta gegenbauri is abundant in the North Atlantic and occurs in frequent blooms [74]. Polar lipids were the dominant lipid classes. The FA composition was largely dominated by EPA (13–14%) and DHA (27–30%).

8 Marine invertebrates 8.1 Sponges Marine sponges are the most primitive multicellular animals and contain many new metabolites, including lipids, in particular glycolipids and phospholipids [91–97]. Thus, sponge lipids are one of the richest source of unusual FA. Sponges are very ancient animals with special structural features in their cell membranes, in particular phospholipid FA and sterols, since sterol– phospholipid interactions are assumed to play a major role in cell membranes [91–93]. Furthermore, sponge classification needs to be supported by chemotaxonomic criteria, in particular regarding FA. Recently, a comprehensive taxonomy was published, which provides the state of the art [16]. Marine invertebrates, e.g. sponges, are filter feeders and consequently they can be associated with microorganisms. Thus, particular FA appear as biomarkers for such organisms. It has been chosen here to focus on the most unusual recent data, such as unsaturated or branched patterns. 8.1.1 ∆5,9 fatty acids Major sponge phospholipid FA include very low amounts of the usual methylene-interrupted PUFA, or none at all, but high levels of very-longchain acids (C23 to C34 , representing up to 80%); these are the so-called demospongic acids. Sponges (Demospongia) are a source of novel FA structures, especially unusual long-chain ∆5,9 FA with no counterpart in the terrestrial world, with sometimes a third double bond or a bromine atom [93, 96–102]. In each set of sponge phospholipids studied, about 50 to 70 FA were

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identified. Some sponges contain up to fifteen ∆5,9 FA in their phospholipids and, in several cases, one of them accounted for more than 50% of the total phospholipid FA mixture [99, 102]. Such NMI FA are showed in Fig. 7. Approximately twenty ∆5,9 unsaturated FA were found in these sponges [102]. It is generally admitted that ∆5,9 demospongic acids are biosynthesized by Demospongiae through short-chain unsaturated FA, mainly from exogenous ∆9-16:1 [93]. The sponge Geodinella robusta was very interesting in that it contained an unusually high level of free FA, mainly with ∆5,9 unsaturation, including the rare anteiso-5,9-24:2 acid (19.5% of the total free FA), and the new iso-5,9-24:2 acid (30%) [103]. Mixtures of these latter compounds showed cytotoxic activity against mouse Ehrlich carcinoma cells and a hemolytic effect on mouse erythrocytes [103]. The FA of total lipids from Halichondria panicea included ∆5,9,19-26 a major component (33%), and six other ∆5,9 FA [104]. Several common PUFA, such as AA, EPA and DHA, occurred in this sponge. The 5,9,23-triacontatrienoic methyl ester, isolated as a natural compound by bioassay-guided fractionation from the marine sponge Chondrilla nucula, is an elastase inhibitor with the potential to be a therapeutic agent in some diseases such as pulmonary emphysema and chronic bronchitis [105]. The 5,9,21-30:3 sponge acid was reported as a DNA topoisomerase inhibitor [106]. Brominated FA from marine invertebrates have also been reviewed [100]. Recently, a comprehensive review of natural halogenated FA included those from marine algae and invertebrates [36]. Cinachyrella sponges from the Red Sea also contained three 6-brominated acids with the new 6-bromo-5,9nonacosadienoic [102]. 6-Bromo-5,9,24-27:3 and 6-bromo-5,9,24-28:3 acids have been isolated from the sponge Xestospongia sp. [107, 108]. It should be noted that the latter FA, and the 6-bromo-5,9-heptacosadienoic acid, displayed moderate activity against murine leukaemia L1210 cells and against human carcinoma KB cells [108]. Nevertheless, it was demonstrated that sponges are not the only source of these unusual ∆5,9 FA since they have been observed in zoanthids and sea anemones [109, 110], gorgonians [100, 111]. Thus, the novel iso-5,9pentadecadienoic acid was isolated from Eunicea succinea and prepared by synthesis [111]. In addition, new 6-bromo-5,9-eicosadienoic acid was identified from an anemone and a zoanthid [110].

Fig. 7 ∆5,9 fatty acids from Cinachyrella sponges

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Fig. 8 Irciniasulfonic acid from Ircinia sp.

A novel FA analogue, named irciniasulfonic acid, has been isolated from the Japanese marine sponge, Ircinia sp. Its structure consists of three different kinds of acids, i.e. common FA, a novel unsaturated branched C10 FA and an isethionic acid. Irciniasulfonic acid and deacyl irciniasulfonic acid reverse multidrug resistance in human carcinoma cells caused by overexpression of membrane glycoprotein [112] (Fig. 8). 8.1.2 Branched fatty acids Branched FA including isoprenoid acids have often been found in marine sponges. The main isoprenoid FA are 4,8,12-trimethyltridecanoic and phytanic acids [95, 96, 101, 102]. As a recent example, phospholipid FA of two sponges of the genus Cinachyrella from the Red Sea were studied and compared with previous results for other Cinachyrella sponges [102]. Five SFA not hitherto found in nature were identified, namely 17-methyl-24:0, 18-methyl24:0, 18-methyl-25:0, 18-methyl-26:0 and 18,24-dimethyl-26:0 acids [102] (Fig. 9). The rare 10,13-dimethyl-14:0 and the new 9,13-dimethyl-14:0 were identified in marine sponges [102, 114]. The occurrence of bacteria in sponges is supported by the huge levels of phosphatidylglycerol and phosphatidylinositol that are characteristic of bacterial membranes [93]. In addition, several branched short-chain FA were identified, which are typically of bacterial origin, such as iso- and anteiso-15:0, iso- and anteiso-17:0, 10-methyl-16:0, 13-methyl-16:0, 10-methyl-18:0, and 11-methyl-18:0 acids. The new branched long-chain acids probably arise from shorter precursors of exogenic origin through a homologation process [102]. Other sponges were found to be rich in such branched FA, including additional branched FA, 3-methyl branched short-chain acids and the new 8,10-dimethyl-16:0 acid [101, 115–117]. The lipids of the sponge Hymeniacidon sanguinea from the Black Sea contained 73 FA, including the novel 13-methyl-20:0, 15-methyl-22:0 and 3,13-dimethyl-

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Fig. 9 New long-chain branched FA from Cinachyrella sp.

14:0 [116]. The compositions of lipids in this sponge collected from two locations with different ecological conditions (Canary Islands and Black Sea) were compared [94, 116]. The significant differences in the structures and relative concentrations of the typically bacterial FA, and the occurrence of cyclopropane-containing FA only in the Canary Islands sponge, suggests that the symbiotic bacteria in the two species are different [116]. A most interesting finding in Callyspongia fallax was a series of iso monoenoic branched-chain C15 -C17 and double bonds at either ∆4, ∆5, ∆6, ∆7, or ∆9 [117]. The lipids of the stromatoporoid Astrosclera willeyana (a “living fossil” sponge) and the demosponge Agelas oroides contained complex isomeric mixtures, at large amounts, of branched FA including iso-/anteisobranched FA and abundant mid-chain branched acids present in the C15 to C25 range. These compounds are most likely derived from specific heterotrophic bacterial symbionts [118]. 8.1.3 Methoxylated fatty acids Methoxylated FA are relatively rare in nature and limited to primitive organisms such as cyanobacteria, bacteria and sponges [66, 101, 117, 122]. The first naturally occurring α-methoxylated FA were found in phospholipids from the sponge Higginsia tethyoides, which contained saturated, monounsaturated and diunsaturated α-methoxylated FA with chain lengths between 19 and 28 carbon atoms [121, 122]. Methoxylated lipids have been recently reviewed with an emphasis on the alkylglycerol ethers and FA bearing the methoxy group in the alkyl chain [119]. In that recent review, 29 αmethoxylated acids were listed [67]. These phospholipid α-methoxylated FA share the common molecular properties of possessing the R configuration at the chiral center [123]. While the first very-long-chain α-methoxylated FA (C19 –C28 ) could have arisen from sponge cells, recent examples of shortchain analogs (C14 -C18 ) are postulated to originate from bacteria in symbiosis with sponges. A novel series of α-methoxylated FA have been reported from Caribbean sponges from the genera Amphimedon, Callyspongia and Sphe-

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Fig. 10 Some α-methoxylated FA identified in Caribbean sponges

ciospongia (Fig. 10) [122, 123]. Various syntheses of methoxy FA have been performed [124]. 8.1.4 Acetylenic fatty acids Acetylenic FA have been found in sponges [125, 126]. Brominated C16 , C18 , and C20 acetylenic FA have been reported from sponges of the genera Xestospongia and Oceanapia [127]. An already known brominated acetylenic acid was isolated from the sponge Xestospongia testudinaria by bioassay fractionation (A1 adenosine receptor affinity), as shown in Fig. 11 [128]. This FA was the active compound. Two novel steryl esters with bromoacetylenic chains that were also isolated were found to be inactive. A C14 acetylenic acid from the sponge Oceania sp. showed significant antimicrobial activity against various bacteria and fungi (Fig. 12a) [129]. Recently, new acetylenic FA from the Steletta sponge species exhibited weak cytotoxicity against a human leukemia cell line [130] (Fig. 12b). The second compound from Stellata is a symmetric dimer of the first, and is the first example of a sponge metabolite possessing an acid anhydride functionality (Fig. 12b). In a recent work, a sulfated C24 acetylenic FA, namely callysponginol sulfate A, was isolated by bioassay-guided fractionation from the Japanese sponge Callyspongia truncata (Fig. 13) [131]. Callysponginol sulfate A is the first example of an acetylenic acid containing a sulfate group from marine organisms. These compounds inhibited membrane type 1 matrix metalloproteinase (MTP1-MMP), one of the key enzymes involved in tumor growth, migration, angiogenesis, invasion and metastasis [131]. Recently, new lysophosphatidylcholines and monoglycerides were reported from Stelletta, which include acetylenic fatty acyl chains and dimethyl branched chains [132].

Fig. 11 Biologically active FA from Xestospongia testudinaria

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Fig. 12 New acetylenic FA from (a) Oceanapia and (b) Stellata

Fig. 13 Callysponginol sulfate A from Callyspongia truncata

Ceramides have received increasing interest because of their various properties, including antifungal and antimicrobial activities. Recently, several new ceramides have been isolated from various marine sponges including Haliclona tenuiraniosa (Fig. 14A) [133, 134]. They contain the usual saturated C20 to C26 fatty acyl chains. Biofouling organisms such as barnacles, mussels and macroalgae cause serious damage to ship’s hulls and aquaculture nets. Thus, it is very important to identify nontoxic alternatives to the organotin compounds currently used. The sponge H. koremella provides a new ceramide with activity as an antifouling substance against macroalgae (Fig. 14B) [133]. This work showed that the length of the acyl residue seems to be important for the antifouling activity of ceramides. Recently, a mixture of ceramides has been isolated from the red alga Ceratodictyon spongiosum containing the symbiotic sponge Sigmadocia symbiotica [135]. Their non-hydroxylated acyl chains ranged from C22 to C24 . In addition, a unique 24-methylenecholesteryl ester (19:0) was characterized in these organisms. Glycosphingolipids (GSL) are ubiquitous membrane constituents in animals that play fundamental roles in major phenomena such as cell–cell recognition and antigenic specificity. GSL from marine sponges possess interesting immunostimulatory and antitumor activities [9–11, 136–141]. Sponges are a very rich source of new glycolipid structures [9–11, 139–141]. Two glycolipids were isolated from the sponge Pseudoceratina sp.: a galactosyl diacyl-

Fig. 14 Ceramides from the sponges Haliclona tenuiraniosa (a) and H. koremella (b)

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glyceryl, and an alkylacylglycerol linked to a five-membered cyclitol, with common fatty acyl chains ranged from 14:0 to 18:0 [142]. Ectyoceramide, the first natural hexofuranosylceramide has been isolated from Ectyoplasia ferox in a pure form, instead of a complex mixture as is usually found [11]. The FA attached to the amino group is a 2-hydroxylated anteiso-16:0. It has been demonstrated that a change in the stereochemistry of the glycosidic linkage from the usual β to the quite rare α configuration can affect their biological activity. Thus, α-glycosyl GSL, such as the natural Agelasphines and their synthetic derivative KRN7000 [143–145], are immunomodulating and antitumor compounds. Not all sponge glycolipids are GSL and a great variety of structures is observed. These compounds stimulated the microtubule polymerization at 10 ◦ C. The investigation of glycolipids from the Caribbean sponges allowed the isolation of Plakosides A and B from Plakortis simplex [10] (Fig. 15), and Plakosides C and D from Ectyoplasia ferox [140] (Fig. 15). These glycolipids are among the most fascinating lipids isolated from marine organisms. They have a unique structure with a prenylated galactose and two cyclopropanyl alkyl chains. The Plakosides had the same α-hydroxylated C22 fatty acid amid with a cyclopropane at the C11–C12 positions. As the sponges are taxonomically distant, it is possible that these unusual glycolipids may originate from bacteria. Plakosides A and B are immunosuppressants that act through a noncytotoxic mechanism [9, 10]. Crasserides, and their minor associated compounds isocrasserides, are widely distributed in marine sponges, and are considered to be glycolipids although their sugar unit is replaced by an unusual five-membered cyclitol [146] (Fig. 16). The cyclitol is linked to the glycerol with an O-3 ether bond. Also linked to the glycerol are an O-1 alkyl group and an O-2 acyl group. Furthermore, various fatty acyl chains are present including mid-chain branched chains. The Italian group has found crasserides in all sponges whose glycolipids have been studied.

Fig. 15 Plakosides A and B from Plakortis simplex, and Plakosides C and D from Ectoplasia ferox

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Fig. 16 An example of an isocrasseride isolated from Plakortis simplex

8.2 Coelenterate – Cnidaria Representatives of the Coelenterate phylum have remarkable peculiarities in their FA composition. Some species contain unusual FA or unusual concentrations of common components. Thus, large amounts of tetracosapolyenoic acids, namely 24:6(n-3) and 24:5(n-6), were found in different orders of the Octacorallia subclass of Anthozoa [147]. Gorgonian corals are of interest since some of these invertebrates are known sources of methyleneinterrupted PUFA, in particular of the (n-3) and (n-6) series [148, 149]. The gorgonian specimens harvested in colder waters contained high amounts of methylene-interrupted PUFA, unlike specimens from warmer waters. This could have been due to the temperature or to the high content of wax esters. Nevertheless, arachidonic acid, a major component in all FA mixtures studied (14–21%), is a well-known precursor of prostanoid compounds. The high levels of tetracosapolyenoic FA in specimens from colder waters were of particular interest: 24:6(n-3) (5.1–5.3%), 24:5(n-6) (8.4–15.8%) and 24:5(n-3) (5.0–5.2%) [149]. An analysis of four gorgonians from the genus Pseudopterogorgia revealed that the main PUFA are 18:3(n-6), 18:4(n-3), AA, DHA, 24:5(n-6) and 24:6(n-3), with the (n-6) PUFA predominating [150]. All five gorgonians of the genus Eunicea presented a similar phospholipid FA composition with unsaturated acyl chains from C18 to C24 , as shown below in Table 1 [151]. 2-Hydroxy long-chain acids also occurred in these gorgonians. Several 2-hydroxy FA have been identified before in marine sponges [152, 153]. The phospholipid FA composition of the Caribbean gorgonians Gorgonia mariae and Gorgonia ventalina (Gorgoniidae) was investigated [154]. This study reports that the main FA were 14:0, 16:0, 18:3(n-6), 18:4(n-3), 18:2(n-6), AA, DHA and 24:5(n-6), as shown in Table 2. In both gorgonians (n-6) PUFA predominated over the (n-3) family. In addition, Table 2 gives data for other gorgonians of the family Gorgoniidae [150].

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Table 1 Principal unsaturated FA in phospholipid from gorgonians of the genus Eunicea [151] Fatty acid

Eunicea sp.

E. fusca

E. laciniata

E. mammosa E. succinea

18:4(n-3) 18:3(n-6) 18:2(n-6) 18:1(n-9) 20:5(n-3) 20:4(n-6) 22:6(n-3) 24:5(n-6) 24:6(n-3)

24.6 18.0 0.7 4.2 1.3 12.4 5.1 3.8 0.6

16.6 15.8 1.5 9.6 1.5 12.4 6.3 1.5 0.4

7.5 17.0 4.9 2.4 3.1 13.5 4.2 2.5 1.4

7.6 9.8 1.3 1.0 1.3 15.7 3.5 14.2 2.9

15.2 10.2 1.5 2.2 1.5 11.5 6.2 3.9 2.7

Several species also contained 2-OH-20:0 and 2-OH-22:0 acids up to 5% of total FA PL. All species contained the new (Z)-7-Me-16:1(n-10) and (E)-7-Me-16:1(n-10) (0.5–2%)

Within the experimental errors, all of these gorgonians share a similar phospholipid FA profile, in as much as: 1) they all biosynthesize, in a similar ratio, the acids 24:5(n-6) and 24:6(n-3), with the former predominating, 2) the (n-6) family of FA predominates over the (n-3) family, and 3) the polyunsaturated FA DHA and AA are key FA in these gorgonians. Several new branched unsaturated FA occurred in gorgonians [149–151]. The New Caledonian gorgonian Rumphella aggregata also contained 24:5(n-6) (11%), NMID FA, and unusual short branched-chain unsaturated acids [155].

Table 2 Main phospholipid unsaturated fatty acids of gorgonians from the family Gorgoniidae Fatty acids

Gorgonia mariae

Gorgonia ventalina

Pseudopterogorgia∗

18:3(n-6) 18:4(n-3) 18:2(n-6) 18:1(n-9) 20:4(n-6) 20:5(n-3) 22:6(n-3) 22:4(n-6) 24:5(n-6) 24:6(n-3)

10.6 10.8 8.6 5.0 10.0 6.4 5.6 2.2 4.0 2.0

15.8 16.4 10.3 4.0 9.4 1.2 8.6 0.3 1.4 0.4

7.3 12.0 4.6 3.0 17.2 0.2 6.1 – 10.2 3.7

∗

average of 4 Pseudopterogorgia species (Z)-7-Me-16:1 n-10 and (E)-7-Me-16:1 n-10 occurred at 0.8–3.5% 2-OH-21:0 and 22:0 acids were identified at < 1%

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An interesting investigation on the FA composition of solar coral Heliopora coerulea (Octocorallia, Helioporacea) supported a chemotaxonomic significance of tetracosapolyenoic acids in Coelenterates [148]. 24:6(n-3), a tetracosahexaenoic acid, was found in Heliopora coerulea [148]. The major FA were 16:0 (40–42%) and 18:3(n-3) (15–16%), as shown below. Table 3 Main fatty acids from total lipids of Heliopora coerulea [147] Fatty acids

% for two samples

Fatty acids

% for two samples

14:0 16:1(n-7) 16:0 18:1(n-9) 18:3(n-6)

4.8–4.8 3.1–3.3 40.9–41.6 3.1–3.5 15.1–15.8

18:4(n-3) 18:0 20:5(n-3) 22:6(n-3) 24:6(n-3)

3.5–3.6 7.3–5.0 5.4–5.5 4.7–5.5 1.7–1.9

Despite a relatively low content of 24:6(n-3) (2% of total FA), it was concluded that PUFA of regular structure with 24 carbon atoms and five to six methylene-interrupted cis-double bonds are typical constituents of representatives of all orders of the Octacorallia, Alcyonaria, Gorgonaria, Helioporida and Pennatularia. Three novel 10-hydroxydocosapolyenoic acids were isolated from deep-water scleratinian corals, as shown below [156].

Fig. 17 10-Hydroxypolyenoic FA from Madrepora oculata

A work has recently been carried out aimed at elucidating the biosynthesis of docosahexaenoic acid in trout liver microsomes [157, 158]. This report conclude that the tetracosahexaenoic acids, such as 24:6(n-3), are intermediates in the biosynthesis of DHA. Therefore, it is very likely that gorgonians utilize a similar biosynthetic route, thus providing yet another interesting system to study this type of biogenesis. However, work still remains to be done on the role of symbiotic zooxanthellae in the production of some of these unusual FA. A survey of lipid and FA composition was made for 15 cnidarians from Okinawa, Japan [159]. Corals having symbiotic Zooxanthellate within their cells contain large amounts of lipid in their tissues (24–26% of dry weight).

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All specimens contained monoalkyldiacylglycerol and were rich in wax esters and triacylglycerol. Palmitic acid was the most abundant FA component of these lipid classes (more than 40% in each class), followed by stearic and oleic acids [159]. Four ceramides with the same hexadecyl acyl chain isolated from a gorgonian exhibited significant human cholesteryl ester transfer protein inhibitory activity [160]. 8.3 Echinodermata Significant amounts of tetracosapolyenoic acids such as 24:6(n-3) were found in different marine organisms, echinoderms (Ophiuroidea and Crinoidea) and coelenterates [147], and in their predators [161]. 24:6(n-3) occurred in symbiotic and non-symbiotic brittlestars [162]. Isolated from the brittlestar Ophiura sarsi [163], this FA had anti-inflammatory and anti-allergic properties similar to those of DHA [164]. Total lipids of this organism contained 14% of 24:6(n-3), 15% of EPA and 2.6% of DHA [163]. Furthermore, high levels of EPA and 24:6(n-3) were observed in phospholipids that accounted for more than 50% of total lipids from O. sarsi. Several novel NMI FA were further identified in O. sarsi [165], namely 7E,13E-20:2, 7E,13E,17Z-20:3 (13% of total FA), 9E,15E,19Z-22:3, and 4Z,9E,15E,19Z-22:4 acids, as shown below.

Fig. 18 New non-methylene-interrrupted FA from Ophiura sarsi

Other NMI FA also occurred in this brittlestar, namely 5,11-20:2, 5,1320:2, 7,13-22:2 and 7,15-22:2. 24:6(n-3) Accounted for about 6%. Structural analyses of these fatty acids were performed after partial hydrogenation with hydrazine sulfate and subsequent isolation of the monoenoate products by argentation thin-layer chromatography, followed by GC/MS analyses of dimethyl disulfide adducts [165]. A comparison of FA in symbiotic and nonsymbiotic brittlestars from Oban Bay, Scotland, was performed in order to look for bacterial signals that might indicate contribution of subcuticular bacteria to their hosts’ diet [162]. Odd-chained and branched FA were present in low amounts but palmitoleic and cis-vaccenic acids, also considered as good

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markers for bacteria, occurred at much higher amounts in symbiotic species studied than in the non-symbiotic species. EPA occurred at high levels in all species studied (11–23%) but DHA was present at low amounts. The unusually concentrations of 24:6(n-3) (up to 15%) may indicate that DHA derived from dietary phytoplankton is being elongated to the former [162]. Furthermore, three other tetracosapolyenoic acids and 26:6(n-3) were found in some species [162]. Three new ceramides from the starfish Acanthaster planci have been reported [166]. Several glycosphingolipids have been identified recently in sea cucumbers possessing α-hydroxylated or non-hydroxylated, saturated or monoenoic fatty acyl chains [167–169]. 8.4 Tunicates Few works have been published on the lipid composition of tunicates [147, 157, 170, 171]. Lipids of edible ascidian Halocynthia roretzi, very popular in Japan and Korea, have also been studied [170]. Several studies of phospholipids of pelagic tunicates (belonging to gelatinous zooplankton) have also been undertaken [171]. PUFA represent the most important class, accounting for around 50%. The tunicates Eudistoma bituminis and Cystodytes violatinctus from the Indian Ocean were investigated for their phospholipid FA content [172]. In both cases, the most abundant FA were the saturated ones (C10 to C18 ). Cystodytes violatinctus contained high amounts of oleic acid (20%). Both E. bituminis and C. violatinctus contained phytanic acid and ∆10 FA, which had not previously been found in such organisms. These tunicates contained only trace amounts of PUFA, which are usually predominant in this phylum [172]. 8.5 Molluscs 8.5.1 Introduction Lipids are a very important food reserve, in particular in the oocytes of molluscs, which assures viability of the larvae. Lipids also provide energy for growth during conditions of limited food supply when carbohydrate levels (the main energetic reserve in molluscs) are low. Phytoplankton represent the largest food source for bivalve molluscs and contain a high proportion of PUFA. The intensive rearing of bivalves still relies on the massive production of unicellular algae especially for growing young spat, which represent the largest biomass in a commercial hatchery. The high cost and unpredictable culture success of algae has inspired the development of artificial diets such

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as microcapsules, mixed diets, yeast based diets, lipid microspheres and liposomes to substitute or supplement live algal diets. The importance of lipids in bivalve nutrition is now well known [173]. The (n-3) PUFA, EPA and DHA, have been reported to be essential for optimal growth for at least some species of juvenile bivalves. The biochemical composition of the intertidal rocky-shore bivalves, e.g. mussels, is greatly affected by periods of air exposure, at which times the bivalves are denied a food source. Consequently, the resulting effect would be similar to starvation [174]. In coastal environments, detritus, bacteria and zooplankton may greatly affect available food composition. It is known that organic detritus areas an energy source for bivalves during periods of scarce primary productivity. Additionally, detrital material is a source of saturated and monounsaturated C14 –C18 FA. A high proportion of SFA, such as 20:0, has been observed in bivalves distributed in environments rich in organic material with an abundant bacterial load [175], compared with those mainly nourished by marine phytoplankton, which are dominated by (n-3) PUFA of 18, 20 and 22 carbons. The lipid composition of molluscs can be affected by external factors, such as fluctuations in the environmental conditions and qualitative and or quantitative changes in food availability, or by internal factors, such as sexual maturation [175]. Recently, some very interesting investigations have dealt with the chemical composition and chemotaxonomic of cardiolipids from some marine bivalves and led to evidence of a tetradocosahexaenoic cardiolipin [176]. 8.5.2 Mussels FA profiles of seeds of the mussel Mytilus galloprovincialis originating from two habitats (rocky shore and subtidal) were compared after transfer to the same habitat (subtidal), in order to study the initial levels of different FA of metabolic importance and their variability [177]. According to previous studies, PUFA were found to be the group with highest percentage (42–49% of total FA), including EPA and DHA as major components [177]. Moreover, these findings concur with recent studies of numerous bivalve species distributed in other regions of Europe and America, such as Argopecten purpuratus [178] and Crassostrea gigas [179]. Additionally, the mussels of subtidal origin presented higher initial levels than the rocky-shore mussels with regard to FA characterized by energetic-type functions, such as 14:0, 16:0 and EPA. High initial levels of some PUFA observed in the subtidal mussels, such as 18:3(n-3), 18:4(n-3) and EPA, are presumably due to the fact that these mussels had greater access to phytoplanktonic food. Initial levels of 14:0, 16:0 and 18:0 in the rocky-shore mussels were significantly greater than in the mussels of subtidal origin. FA characterized by structural-type functions, e.g. 18:0, DHA and NMID FA with 20 and 22 carbons, in rocky-shore mussels

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presented higher levels than those of the subtidal mussels. These NMID FA have been observed in greater proportions in phospholipids, thus implying a structural-type function [177]. In addition, it is also known that these NMI FA are distributed in greater quantities in the organs more exposed to the immediate environment, such as the gills, mantle and foot. In another study, minor NMI FA were characterized in Mytilus galloprovincialis by GC/MS of their 2-alkenyl4,4-dimethyloxazoline derivatives, namely 7,13–16:2, 5,11–20:2, 5,13–20:2, 7,15–22:2, and the new trienoic FA 5,11,14-20:3 and 7,13,16-22:3 [180]. This discovery supports a biosynthetic route implying the desaturation and subsequent elongation of 20:2(n-6) [180]. Lipid, FA and sterol composition of the New Zealand green-lipped mussel (Perna canaliculus) and the Tasmanian blue mussel (Mytilus edulis) have also been reported [181]. 8.5.3 Oysters Seasonal variations of lipid classes and FA in the flat oyster Ostrea edulis have been studied [182]. The dynamics of FA in the larval development, metamorphosis and post-metamorphosis of this oyster have been investigated [183]. The lipid composition of Crassostrea gigas was analysed during the reproductive phase in natural as well as under artificial conditions [179]. A specific accumulation of DHA and EPA in the polar lipids was observed under both conditioning diets. The proportions of DHA and EPA from neutral and polar lipids of oysters conditioned artificially were significantly lower than of those that were naturally conditioned. A useful comparison of the lipid class and FA composition between a reproductive cycle in nature and a standard hatchery conditioning of the Pacific oyster Crassostrea gigas was performed [179]. 8.5.4 Patella The effects of season and spatial distribution on the FA composition of four Patella species’ gonads and soft body tissue were evaluated [185]. The results show that the quantitatively most important FA were 14:0, 16:0 and 18:0; the MUFA 18:1(n-7), 18:1(n-9), 16:1(n-7) and 20:1(n-9), EPA and AA. P. depressa and P. ulyssiponensis soft-body FA profiles revealed significant differences between sexes, with males showing significantly higher percentages of PUFA, mainly EPA and AA, while in females significantly higher proportions of MUFA were found. The fatty-acid composition of P. depressa gonads revealed significant differences between sexes. Males showed a significantly higher percentage of PUFA from the (n-3) and (n-6) series (AA and EPA), while females were seen to have higher proportions of SFA and MUFA [185].

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8.5.5 Clams Recent studies on clam lipids were concerned with their aquaculture. Thus, the possible use of emulsions rich in EPA and DHA as an artificial lipid supplement to live algae was investigated for seed of the Manila clam Tapes philippinarum [173]. In addition, the influence of the lipid composition of microalgal diets and cornstarch on the lipid classes and FA of the Ruditapes decussatus spat was studied [186]. This clam species is of commercial interest in Spanish aquaculture. In order to increase its production, experiments have been carried out with alternative foods to live microalgae, such as freeze-dried microalgae or cornstarch. The main FA present in the spat of R. decussatus were 16:0, 18:1(n-9) and DHA, followed by lower contents of 18:4(n-3) and 18:1(n-7). The essential FA EPA is present in small amounts. The content of (n-3) PUFA, (n-9), (n-6), and 20:2 and 22:2 NMID FA differed significantly according to the diet supplied. Spat fed on a microalgal diet show the significantly highest content in (n-3) PUFA and (n-9) FA [186]. 8.5.6 Scallops Overfishing has often resulted in a decline of natural beds of Pecten maximus in several areas. Thus, several experiments have been performed on P. maximus culture, growth and reproduction to improve the biological knowledge of this species [187, 188]. Lipids are a very important food reserve in the oocytes, which assures viability of the larvae. It has been shown that success of Pecten maximus hatching is related to the lipid status of the eggs when spawned. Lipids in the female gonad were analysed for lipid class composition and FA composition of TAG and phospholipids, and their variations in relation to gametogenic cycle were studied [184–188]. PUFA were more abundant in the TAG and the series (n-3) was clearly predominant. The principal (n-3) FA (mainly in phospholipids) showed a seasonal variation clearly related to the reproductive cycle [188]. Soudant et al [184] have analysed the composition of polar lipid classes in male gonads of Pecten maximus and the effect of nutrition. Seasonal digestive-gland dynamics of the scallop Pecten maximus have been reported [189]. The distribution of lipids and FA in different organs of the Chilean scallop Argopecten purpuratus broodstock, namely female and male gonads, digestive gland, gills, mantle and adductor muscle, have been investigated [178]. The highest level of (n-3) PUFA (mainly EPA and DHA) was found in the adductor. The major FA in all parts studied were 16:0, EPA (8.1–20.3%) and DHA 9.2–25.6%). Similarities between the FA composition of the triglyceride fraction of the female gonad and the digestive gland (e.g. the high level of 14:0 and 18:4(n-3)) indicated the transfer of lipids from the lipid-rich digestive

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gland to the female gonad. A special feature of the gills and mantle was the presence of high levels of plasmalogens (phosphoglycerides with 1-alkenyl chains) recognized by the presence of dimethylacetals, which are formed simultaneously with FAME by acid-catalyzed transmethylation [178]. In another investigation, dietary supplementation with lipid emulsions during broodstock conditioning of A. purpuratus was used to manipulate the fatty-acid composition of the eggs [187, 190]. The scallops were fed a mixed algal diet either alone or supplemented with an emulsion rich in ethyl esters of DHA or EPA. Lipid supplementation resulted in a significant increase of the total lipid content of the eggs. The EPA and DHA levels in the total and neutral lipids of eggs from broodstock supplemented with diets including the corresponding emulsions were significantly higher than in eggs from scallops fed solely algae [190, 191]. NMID FA, which are not present in the phytoplankton, have been reported in many species of molluscs. Thus, they are presumably synthesized by molluscs. Interestingly, the identification and occurrence of a novel cis-4,7,10,trans-13-docosatetraenoic acid in the female gonads of the scallop Pecten maximus was described [192]. Lipid deterioration during frozen storage at – 20 ◦ C of the adductor muscle, the major edible part of the giant scallop, was examined by determination of fatty chain compositions in the sn-1 and/or sn-2 positions of ether and ester glycerophospholipids [193]. During storage, the contents of total lipid and polar lipid decreased but that of non-polar lipid increased. The percentages of PUFA such as EPA and DHA in the total lipid and polar lipid fractions decreased during storage, but those of the PUFA in the non-polar lipid increased. Changes during storage in alkenyl and alkyl chain compositions of ether glycerophospholipids and in fatty acyl chain compositions of ether glycerophospholipids were determined [193]. The 20:2 and 22:2 NMID FA also occurred. 8.5.7 Squids Biochemical composition changes and FA in several tissues of the squid Illex argentinus from the South Atlantic Ocean at different sexual stages were studied. All tissues contained high concentrations of PUFA followed by SFA and MUFA, e.g.16:0, 18:0, 18:1(n-9), 20:1, 22:1, EPA and DHA. The lipids also contained unusually high levels of FA of the linoleic family (2–3%). The digestive gland of the squid is rich in polyenoic FA, EPA and DHA. This tissue could be a cheap raw material, presently discarded, for production of PUFA [194]. The lipid content and composition of phosphatidylcholine and FA were determined in tissues of 70 species of teleosts, and in muscle of six species of squid. Amount and composition of diacyl glyceryl ethers were re-

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cently analysed in various tissue lipids of the deep-sea squid Berryteuthis magister [195].

9 Crustacea The mud crab Scylla serrata is a commercially important species in the IndoPacific region and has been cited as a target species for a stock-enhancement program in Japan and also as a target species for aquaculture in many Asian countries [196]. A recent study evaluated the requirements of linoleic acid (18:2(n-6)), linolenic acid (18:3(n-3)), EPA and DHA during rotifer, as well as Artemia feeding on survival and larval development of mud-crab larvae [196]. Decreased natural seed availability and the low survival rate in crab hatcheries [197–199] have been major problems in increasing aquaculture production. Moreover, Takeuchi et al [197] described the requirement of EPA and DHA for larval development, where EPA is effective in maintaining survival while DHA plays an important role in accelerating the intermolt period and produces a wider carapace width in swimming crab larvae. A previous study showed that the Artemia feeding schedule (in combination with rotifers) for larval mud crab and their essential FA composition affected the survival of larvae [199–201]. If the conditions of thermal adaptation are well documented in fish, little information is available for marine invertebrates. A comparative study of the phospholipid FA compositions was conducted with the Baltic Sea amphipod crustacean Gammarus spp. collected from different thermal environments [202]. It was reported that environmental temperature had little effect on FA composition. In fact, Gammarus was shown to use the same strategy to control membrane fluidity in the cold as fish species investigated so far [203], namely the crustacean accumulates sn-1 monoenoic and sn-2 polyenoic phospholipid FA at reduced temperatures.

10 Polyunsaturated FA (n-3) of commercial interest 10.1 Introduction An “essential” nutrient is one that is needed for normal development and function of mammalian cells throughout the life cycle. In its active or precursor form there is a minimum amount of such a nutrient that must regularly be provided in the diet. This dietary requirement generally varies with species,

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gender, age and the presence of physiological and pathological challenges (pregnancy, lactation, infancy, aging, infection, disease, etc.). The term “essential fatty acid” is ambiguous and inappropriately inclusive or exclusive of many polyunsaturated FA. When applied most rigidly to linoleate and a linolenate, this term excludes the now well-accepted but conditional dietary need for two long-chain polyunsaturates (arachidonate and docosahexaenoate) during infancy. The metabolism of essential and nonessential FA is clearly much more interconnected than previously understood. Replacing the term “essential fatty acid” by existing but less-biased terminology, i.e. polyunsaturates, (n-3) or (n-6) polyunsaturates, or naming the individual fatty acid(s) in question, would improve clarity and would potentially promote broader exploration of the functional and health attributes of polyunsaturated FA [204]. 10.2 Health benefits Initially, it should be noted that the chemical form of the PUFA supplementation used for clinical assays and up to the final consumers will not be detailed. Indeed, sometimes they take TAG forms while occasionally they are ethyl esters or free fatty acids, depending on the target. Polyunsaturated FA (PUFA) are essential components in higher eukaryotes that confer fluidity, flexibility and selective permeability to cellular membranes. PUFA affect many cellular and physiological processes in both plants and animals, including cold adaptation and survival [205, 206], modulation of ion channels [207, 208], endocytosis/exocytosis [209], pollen formation, pathogen defense, chloroplast development in plants [210], and activities of membrane-associated enzymes that are sensitive to the biophysical properties of lipid membranes [211–213]. In mammals, metabolism of LC-PUFA by oxygenases yields a range of important short-lived molecules (generically known as the eicosanoids), such as prostaglandins, leukotrienes and thromboxanes (Fig. 19). These resulting metabolites bind to specific G-protein-coupled receptors and signal cellular responses and modulate many biological processes (see below). Because the production of various classes of these molecules depends in part upon the availability of their PUFA precursors in membrane phospholipids, modulation of PUFA is a potential target of pharmaceuticals and nutraceuticals [213, 214]. The (n-3) LC-PUFA, particularly EPA and DHA, are thought to display a variety of beneficial effects in areas ranging from foetal development to cancer prevention [215]. Some of those health effects are presented below.

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10.2.1 Heart health It is well established that populations with a high consumption of oily fish have a lower incidence of heart disease and several studies have confirmed that EPA and DHA are the protective components [216–224]. Recent research concludes that perhaps the most important effect of (n-3) LC-PUFA, when it comes to preventing cardiovascular disease, is their ability to stabilize atherosclerotic plaque by reducing the infiltration of inflammatory and immune cells (lymphocytes and macrophages) into the plaque [225]. This could explain the reduction in fatal and nonfatal heart attacks and strokes associated with an increased intake of fish oils [226]. In addition, the antiinflammatory effect of DHA by reducing the C-reactive protein (CRP) level in blood may decrease the risk of coronary heart disease such as atherosclerosis [227]. Several large clinical trials have confirmed the ability of fish oils to prevent sudden cardiac death in both presumably healthy subjects as well as in patients having suffered a heart attack [228–231]. Although most research so far has focused on the effect of (n-3) on life-threatening ventricular arrhythmias it is likely than many of the findings may also be applicable to atrial fibrillation [232]. It has also been shown that a high (n-3) content of blood cells and serum cholesterol esters is associated with increased heart rate variability and leads to a decreased risk of cardiac disease and a longer lifespan [233]. In addition, EPA and DHA seem to reduce the mortality among not only patients who have survived a first heart attack [234–236] but also among old people [237, 238]. Numerous studies have confirmed that (n-3) LC-PUFA included in fish oils significantly combat hypertension by a notable reduction of blood pressure and benefit heart transplant patients [239–247]. EPA and DHA are also effective in lowering the blood level of triglycerides [248, 249], in improving large artery dilation in patients with high cholesterol levels [250], and they possess antithrombotic effect [251]. Such positive actions also contribute to reduce the risk of cardiovascular disease. 10.2.2 Cancer Several studies have shown the effect of LC-PUFA against cancer such as an inverse relationship between blood levels of EPA and DHA and the risk of prostate cancer [252, 253] or fish and fish oil consumption and adenocarcinomas [254]. In addition, (n-3) PUFA can also act positively against cancer effects like cachexia (abnormal weight loss) or survival rate in end-stage cancer [255, 256].

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10.2.3 Arthritis The (n-3) PUFA are also known to decrease rheumatoid arthritis; notably by lowering interleukin-1beta production which results in a significant reduction in morning stiffness and the number of painful joints [257–261]. 10.2.4 Psoriasis Psoriasis is a fairly common skin disease characterized by high concentrations of AA in the plaques and profound changes in the metabolism of eicosanoids leading to an increase in proinflammatory agents. Some studies have shown that n-3 PUFA, notably EPA, can counteract the formation of these proinflammatory agents and that oral supplementation with fish oils benefit psoriasis patients [262, 263]. 10.2.5 Lung disease A few years ago it was shown that children who regularly eat fresh, oily fish have a four times lower risk of developing asthma than children who rarely eat such fish. EPA was suspected to be responsible by reducing airway inflammation and responsiveness. Later, studies on supplementation by (n-3) LC-PUFA have confirmed their benefit in the reduction of breathing difficulties and other symptoms in asthma patients. More recently, it has been demonstrated that those PUFA are also beneficial in the treatment of other lung diseases such as cystic fibrosis and emphysema [264–267]. 10.2.6 Attention-deficit disorder Attention-deficit hyperactivity disorder (ADHD) is characterized by hyperactivity, emotional instability, poor coordination, short attention span, poor concentration, impulsiveness, and learning disorders. It is very common among school-age children with an incidence of 4–20%. Initial studies have linked ADHD to a deficiency of certain long-chain FA, notably AA, EPA and DHA [268]. More recently Burgess et al. [269] have found that children with ADHD were breastfed less often as infants than children without ADHD (breast milk is an excellent source of DHA). In addition to ADHD, other disorders such as dyslexia (difficulties in learning to read and write) and dyspraxia (problems with coordination and muscle control) also have deficiency in LC-PUFA as a common denominator that may be avoided by oral supplementation of concentrated fish oils [270].

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10.2.7 Mental health Several epidemiological studies have shown that a high dietary intake of linoleic acid and a low intake of EPA and DHA are associated with cognitive impairment and an increased risk of dementia. In a recent study, it was that EPA and especially DHA help keep the membranes of brain cells more fluid while saturated and (n-6) FA tend to “harden” them. Authors believe this and the anti-inflammatory effects of EPA and DHA are what help preserve cognitive function [271]. In addition to preventing dementia, (n-3) PUFA help in combating depression, schizophrenia, Alzheimer’s disease and other mental illnesses [272–283]. 10.2.8 Pregnancy and infancy An adequate intake of DHA and EPA is particularly important during pregnancy and lactation. During this time the mother must supply all the baby’s needs for DHA and EPA because it is unable to synthesize these essential FA itself. DHA makes up 15–20% of the cerebral cortex (a normal adult human brain contains more than 20 g of DHA) and 30–60% of the retina (it is also concentrated in the testes and sperm), so it is absolutely necessary for normal development of the foetus and baby, which implies optimal levels in pregnant and lactating mothers. There is some evidence that an insufficient intake of (n-3) FA may increase the risk of premature birth and an abnormally low birth weight. There is also emerging evidence that low levels of (n-3) acids are associated with hyperactivity in children [268, 284–292]. The constant drain on the mother’s DHA reserves can easily lead to a deficiency that may be linked to preeclampsia (pregnancy-related high blood pressure) and postpartum depression [288–290] 10.3 Nutrition: importance of the ratio of (n-6) and (n-3) essential FA There are good fats and there are bad fats. Artificially produced trans-FA are bad in any amount and saturated fats from animal products should be kept to a minimum. The best fats, or oils rather since they are liquid at room temperature, are those that contain the essential FA that are so named because without them we die (see chapter introduction). Essential FA are polyunsaturated and grouped into two families, the (n-6) EFA and the (n-3) EFA. Seemingly minor differences in their molecular structure make the two EFA families act very differently in the body. 18:2(n-6) and 18:3(n-3) are not interconvertible and compete for the rate-limiting 6-desaturase in the synthesis of LC-PUFA (see biosynthesis). AA and EPA are the parent compounds

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for the production of eicosanoids with opposite properties, see Fig. 19. Many scientists believe that a major reason for many diseases (see below) is the profound imbalance between our intake of (n-6) and (n-3) FA. Our ancestors evolved on a diet with a ratio (n-6)/(n-3) of about 1 : 1. A massive change in dietary habits over the last few centuries (modern agriculture) has changed this ratio to something closer to 20:11 [293–295]. An increase in the dietary intake of (n-6) EFA changes the physiological state to a prothrombotic, proconstrictive, and proinflammatory state. Many of the chronic conditions, cardiovascular disease, diabetes, cancer, obesity, autoimmune diseases, rheumatoid arthritis, asthma and depression, are associated with increased production of thromboxane A2 (TXA2 ), leukotriene B4 (LTB4 ), IL-1β, IL-6, tumor necrosis factor (TNF), and C-reactive protein [295]. All these factors increase with (n-6) fatty-acid intake and decrease with (n-3) fatty-acid intake, whether 18:3(n-3), 20:5(n-3) or 22:6(n-3). EPA and DHA are more potent2 , and most studies have been carried out using EPA and DHA (see above). The optimal dose or ratio of (n-6)/(n-3) varies from 1/1 to 4/1 depending on the disease under consideration. In the secondary prevention of cardiovascular disease, a ratio of 4/1 was associated with a 70% decrease in total mortality [296]. A ratio of 2.5/1 reduced rectal cell proliferation in patients with colorectal cancer, whereas a ratio of 4/1 with the same amount of (n-3) PUFA had no effect [297]. The lower (n-6)/(n-3) ratio in women with breast cancer was associated with decreased risk [298]. A ratio of 2–3/1 suppressed inflammation in patients with rheumatoid arthritis, and a ratio of 5/1 had a beneficial effect on patients with asthma, whereas a ratio of 10/1 had adverse consequences [265, 299]. These studies indicate that the optimal ratio may vary with the disease under consideration. This is consistent with the fact that chronic diseases are multigenic and multifactorial. Therefore it appears important to restore the balance between (n-6) and (n-3) for homeostasis and normal development. On this basis and by recognizing the unique benefits of EPA and DHA and the serious consequences of a deficiency, recommendations for daily intake of (n-3) PUFA has been published by several international scientific authorities [300–303]. 10.4 Routes for biosynthesis Despite extensive screenings, no angiosperm plants have been detected that accumulate in reserve triacylglycerols or in membrane lipids cispolyunsaturated FA with carbon chains longer than C18 [305]. However, it 1

Lipid consumption in Europe nowadays: vegetal (65%) > land animals (17%) > butter (16%) > marine animals (2%). 2 Alpha-linolenic acid can be converted to EPA and DHA in the body, but the conversion is quite inefficient, especially in older people [294].

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Fig. 19 Biosynthesis of eicosanoids from the essential FA [304]. Abbreviations: PL, phospholipids; PLA2, phospholipase A2; COX, cyclooxygenase; LOX, lipoxygenase; PG, prostaglandins; TBX, thromboxanes; HETE, hydroxy-eicosatetraenoic acids; HPETE, hydroperoxyeicosatetraenoic acids; LT, leukotrienes. Dietary lipids provide EFA and preformed substrates for the COX/LOX pathways. Dietary FA such as 20:3(n-6), 20:4(n-6), and 20:5(n-3) are direct precursors, while 18:2(n-6) and 18:3(n-3) must be elongated and desaturated prior to their conversions to eicosanoids

is well known that polyunsaturated FA have crucial roles in membrane biology and signalling processes in most living organisms (see the section on health benefits). As indicated by Abbadi et al. [306], the daily requirement of such PUFA may vary and depend on the availability of linoleic and linolenic acid for conversion by elongation and desaturation, but a more regular consumption and an accordingly sustainable source of PUFA would be highly desirable. At present, these FA enter the human diet mainly in the form of marine and freshwater fish. But in view of the increasing human population, the overfishing of marine resources, the dependence of fish farming on PUFA from fish oil and the environmental impact of aquaculture systems, neither farmed nor captured fish can be considered as a sustainable source of PUFA (see section on market) [307]. Thus alternatives have to be found such as new sources or by modifying existing ones by genetic approaches. For example, transgenic oilseeds could be a way out of the forthcoming shortage, particularly in view of the fact that a low percentage of PUFA in daily-consumed plant oils would satisfy nutritional requirements [306]. So, over the last few years, the biosynthetic LC-PUFA pathway has been the subject of much interest and it is only recently that molecular genetic approaches have allowed detailed studies of the enzymes involved in their

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Fig. 20 Pathways of very-long-chain polyunsaturated synthesis in different organisms. Major products are indicated in ovals while enzymes are boxed. The anaerobic route (a) makes use a polyketide-like system (PKS), whereas aerobic routes (b, c) use different enzymes (desaturases and elongases). If routes (b, c) are started with linoleic acid (9,12–18:2), arachidonic acid (AA) = 5,8,11,14–20:4 is obtained. Route (b) is a direct pathway that may operate in marine primary producers and initiate the food chain of “oceanic” polyunsaturated FA ending in large carnivorous fish. Route (c) represents the Sprecher [22] pathway as typical for mammalian cells. Mammals lacks ∆12 and n3 desaturase activities and obtain linoleic acid and α-linolenic acid (9,12,15:18:3) from their diets. In this route, the synthesis of DHA is now known to consist of two succeeding elongation cycles, a ∆6 desaturation and a β-oxidation chain-shortening

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synthesis. Thus, at this time, three distinct routes for LC-PUFA biosynthesis have been identified; two are aerobic and one is anaerobic (Fig. 20). 10.4.1 Aerobic pathways In these routes, LC-PUFA biosynthesis is catalysed by sequential desaturation and fatty acyl elongation reactions. Known pathways involve the processing of the saturated 16:0 or 18:0 products of fatty acid synthase (FA) by elongation and aerobic desaturation reactions. The desaturase enzymes insert double bonds at specific carbon atoms in the fatty acid chain and the fatty acid elongation system elongates the precursors in two-carbon increments [308, 309]. Significant progress has been made in the identification of the enzymes required for PUFA synthesis; in particular, the fatty acid desaturases which are central to this pathway have now all been identified [308, 310]. The most relevant desaturases required for PUFA biosynthesis are the so-called “frontend” desaturases [311] that introduce a new double bond between an existing one and the carboxyl end of the acyl group. These “front-end” desaturases are all members of the cytochrome b5 fusion desaturase superfamily, since they contain an N-terminal domain that is orthologous to the microsomal cytochrome b5 [312]. Interestingly, there is a division of labor in higher eukaryotes for the synthesis of 20:4(n-6), 20:5(n-3) and other PUFA. Angiosperm plants convert oleic acid (18:1) to linoleic acid (18:2) and linolenic acid (18:3(n-3)), which are essential FA for mammals as substrates for the synthesis of C20, but these plants are unable to elongate the FA further. Mammals also convert 18:0 to 18:1(n-9) using a membrane-bound 18:0-CoA desaturase, however, they lack both ∆12 and (n-3) desaturase activities. Therefore, they require 18:2(n-6) and 18:3(n-3), the essential FA (EFA), in their diet [213]. These EFA are converted to long-chain PUFA via a series of desaturation and elongation reactions in the endoplasmic reticulum (ER) [308]. The ∆6 desaturase uses 18:2(n-6) and 18:3(n-3) as a substrates and inserts a double bond to produce 18:3ω6 and 18:4ω3. PUFA with a double bond at ∆6 are substrates for the elongation machinery, which uses malonyl-CoA to add two carbons to the C-terminal end of the FA, producing 20:3(n-6) and 20:4(n-3). These FA are substrates for a ∆5 desaturase, which produces 20:4(n-6) and 20:5(n-3) [213]. Those pathways for the synthesis of arachidonic acid (AA) and eicosapentaenoic acid (EPA) have been characterized biochemically and are supported by the recent cloning and characterization of desaturase and elongase genes. In addition, some notable variations exist such as, for example, the freeliving nematode Caenorhabditis elegans [313–316], the fungus Mortierella alpina [317] the moss Physcomitrella patens [318] the red algae Porphyridium cruentum [319], the freshwater protist Euglena [320] and the microalga

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Isochrysis galbana [321]. All these organisms posses particular enzymatic pathways. At this stage of LC-PUFA biosynthesis a split occur. The force of logic suggested that the steps succeeding AA and EPA synthesis would be another two-carbon elongation step and countervailing desaturation by ∆4 desaturase to produce the C22 PUFA (elongation of EPA and AA to C22:5(n-3) and DTA, respectively, and the desaturation of the latter to form DHA and DPA). The cloning of a ∆4 desaturase from the DHA-producing marine protist, Thraustochytrium sp., suggests that this pathway (route b, Fig. 22) is valid in some organisms [322]. However, mammals lack ∆4 desaturase activity, and evidence has accumulated for an alternative pathway for C22 PUFA biosynthesis. The sequence involving ∆4 desaturase is the simplest one. It operates in various unicellular, eucaryotic algae belonging to different systematic groups and contributing to marine primary production. These and other photosynthetically active organisms are considered to be the primary sources of EPA and DHA entering marine feeding webs with large carnivorous fish and finally humans at the end [306, 307]. The other route, the so-called Sprecher pathway (route c, Fig. 20), remained controversial but recent experiments have indicated that it is the predominant route to DHA in mammals [22, 23, 323–328]. It now seems clear that the human C22 PUFA synthesis pathway consists of two succeeding elongation cycles (leading to 24:5(n-3)) followed by a ∆6 desaturation, all of which occur in the endoplasmic reticulum. After transfer of the fatty acid to peroxisomes, there is a specific β-oxidation chain-shortening to the C22 product. By this route, the synthesis of DHA from acetyl–coenzyme A (acetyl-CoA) requires approximately 30 distinct enzyme activities and nearly 70 reactions, including the four repetitive steps of the fatty acid synthesis cycle [329]. Over the past few years, sequences encoding virtually all the enzyme activities involved in microsomal PUFA biosynthesis have been isolated, identified, and expressed in a variety of heterologous hosts (Table 4) [330]. Results from these studies help to increase our understanding of the biochemistry of elongases and desaturases and the regulation of PUFA biosynthesis. Then, the next challenge will be the selection of a set of suitable copies of the available genes, which after transformation and expression in a heterologous host (such as oilseed crop) should result in a cooperating ensemble of enzymes and production of PUFA. 10.4.2 Anaerobic pathway The diversity of PUFA synthesis described above relies on variations on desaturase and elongase biochemistry. This aerobic biosynthetic pathway was thought to be taxonomically conserved, but nature has also solved the problem of PUFA synthesis using a fundamentally different anaerobic path-

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Table 4 Origin of presently available genes for cDNAs encoding desaturases and elongases involved in the biosynthesis of LC-PUFA. The encoded enzymes have been characterized by functional expression. The numerous sequences published for the ubiquitous ∆9-, ∆12- and ∆15-desaturases are not included [308] Enzyme

Organism

Reference

∆4 desaturase ∆5 desaturase

Thraustochytrium sp Homo sapiens Caenorhabditis elegans Mortierella alpina Homo sapiens Caenorhabditis elegans Borago officinalis Ceratodon purpureus Physcomitrella patens Mortierella alpina Euglena gracilis Caenorhabditis elegans Caenorhabditis elegans Mortierella alpina Physcomitrella patens

[322] [331, 332] [333, 334] [335, 336] [337] [338] [339] [340] [341] [342, 343] [320] [344] [314] [317] [345]

∆6 desaturase

∆8 desaturase (n-3) desaturase ∆6 elongase

way [329]. Indeed, numerous PUFA-producing bacterial strains are capable of producing PUFA under strictly anaerobic conditions, thus precluding the participation of an oxygen-dependent mechanism. The system involved here does not require the multiple desaturase and elongase enzymes outlined above, but instead uses a polyketide synthase-like (PKS) gene cluster, found in both prokaryotic and eukaryotic marine microbes, to synthesize PUFA [214]. Several marine bacteria contain EPA and DHA at levels as high as 25% of total membrane FA [346]. A genomic library prepared from one of these marine bacteria (Shewanella sp. Strain SCRC2738) was used to identify a 38 kb DNA fragment that resulted in EPA synthesis when expressed in E. coli. [347]. Experimental results [329, 348–350] indicated that these genes expressed in E. coli encode a protein complex that is capable of EPA synthesis without any reliance on enzymes of the E. coli FA or any long-chain intermediate such as 16:0-ACP. Apparently, the genes encode a PKS that acts independently of FA elongase and desaturase activities to synthesize EPA directly [213]. Genes with high homology to the Shewanella gene cluster have been identified in Photobacterium profundum [351] and in Moritella marina, which accumulates DHA rather than EPA [352]. Thus it is likely that the PKS pathway for PUFA synthesis is widespread in marine bacteria [213]. The thraustochytrid Schizochytrium sp accumulates large quantities of triacylglycerol rich in DHA (see chapter on thraustochytrids). Biochemical experiments have revealed the involvement of a PKS protein com-

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plex [329], which was further confirmed by molecular genetic analysis. The Schizochytrium genome encodes three proteins with domains highly similar to those encoded by genes from Shewanella, raising the possibility that the PUFA PKS has undergone lateral gene transfer [213]. Several “front-end” PUFA aerobic desaturases from Thraustochytrium have been identified recently [322, 353], but the prevalence of this newly discovered PKS-like biosynthetic pathway is not known. Because Schizochytrium is also a member of the Thraustochytriidae, it is perhaps surprising to find these two distinct biosynthetic pathways represented in the same family, but molecular characterization of PUFA biosynthesis in the Thraustochytriidae might provide insights into the evolution of this important pathway. The primary structure of Shewanella PKS (and its relatives) does not conform to any of the previously described classes of PKS proteins. Instead, it suggests the assembly of several multifunctional proteins into a complex. PUFA PKS carry out some of the same reactions as FA and use the same small protein (or domain), acyl carrier protein (ACP), as a covalent attachment site for the growing carbon chain. However, in these enzyme systems, the complete cycle of reduction, dehydration, and reduction seen in FA is often abbreviated, so that a highly derivatized carbon chain is produced, typically containing many keto and hydroxy groups as well as carbon–carbon double bonds in the trans configuration (Fig. 21). The linear products of PKSs are often cyclized to form complex biochemicals that include antibiotics, aflatoxins, and many other secondary products [329]. Since the double bond on the PUFA molecules were formed by dehydration and isomerization of keto groups in cycles of polyketide-forming chain elongation, reaction to the C20 PUFA may not be the direct precursor of C22 PUFA [354]. The relative simplicity of this PKS-like system makes it attractive in terms of transgenic production of LC-PUFA. For example, introducing and regulating the three Schizochytrium PKS-like open reading frames in a transgenic plant is relatively simple compared with the more than five desaturase and elongase genes required for the aerobic pathway [306]. In addition, the identification of new (PUFA-specific) PKS activities such as double-bond isomerization might help in the bioengineering of additional families of polyketide antibiotics [214]. Furthermore, our improved knowledge of PUFA synthesis in Shewanella, Schizochytrium and their relatives has implications for understanding foodweb dynamics in marine ecosystems. Because these organisms are significant primary producers of 20- and 22-carbon PUFA in cold-water oceans [346], the PKS pathway may be an important source of PUFA for fish and mammals and thus also for human diets. The importance of 20:5 in food-web dynamics of freshwater ecosystems has recently been discussed [355]. Finally, the identification of these PKS systems in ancient lineages raises intriguing questions about the evolutionary relationship of this newly discovered pathway to 20:5

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Fig. 21 Generalized scheme for the processive synthesis of polyunsaturated FA (PUFA) by a polyketide synthase (PKS) system [214]. In the case of PUFA, it is envisaged that a primer molecule (in the form of acetyl-CoA) undergoes several rounds of sequential reactions (keto-synthase, keto-reductase, dehydratase and enoyl reductase), resulting in repeated synthesis and fatty acyl chain (esterified to the acyl carrier protein) elongation by two carbons per cycle. Because PUFA contain methylene-interrupted double bonds (i.e. at the third carbon), it is likely that a dehydratase (FabA-like) module in the PKS also simultaneously carries out a trans–cis isomerization to generate this configuration

and 22:6 FA relative to the enzymatically more complex desaturase/elongase route found in higher eukaryotes [213]. 10.5 Some promising sources of marine LC-PUFA 10.5.1 PUFA from nonphotosynthetic microorganisms If nowadays conventional fish oils are the main industrial sources of PUFA they may be not suitable to meet the increasing markets, notably for DHA owing to their limited supply (see section on sources and market), lower content of DHA in comparison to that of EPA, and peculiar taste and odour. Thus, the production of (n-3) PUFA by microbial fermentation of oleaginous microorganisms has attracted considerable attention in relation to industrial application of single-cell oil (SCO) [354, 356, 357]. For instance, a range of autotrophic and heterotrophic microbes has been assessed for potential commercial sources of EPA and DHA by various workers including Barclay et al. [358], Lewis et al. [359] and Vazhappilly and Chen [360], and the results

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of many studies in this area have been reviewed by Singh and Ward [361] and Ratledge [362]. In comparison to autotrophic microorganisms, the de novo synthesis of (n-3) and (n-6) PUFA by heterotrophic microorganisms may provide an easier and less expensive means of producing PUFA-rich biomass and oils [363]. Thus, Ratledge [362] considered that, despite improvements in the efficiency of photobioreactors, it is doubtful whether the growth of microalgae in bioreactors could be scaled up to satisfy even a modest demand for SCO rich in (n-3) PUFA, and suggested that heterotrophic microbes might be a more productive source. In recent years, interest in the use of microheterotrophs as a source of PUFA has increased [364]. Microheterotrophs do not require some of the elements necessary for the culture of autotrophs (e.g., light, carbon dioxide), and some see them as a potential alternative to traditional commercial sources of PUFA. Arachidonic acid has been produced in quantity by some fungi [365, 366]. Certain bacteria have been shown to produce EPA and DHA [367, 368]. The recognized need in aquaculture for alternative sources of PUFA for feeding both larvae and adults has seen PUFA-producing bacteria successfully demonstrated as a means to enrich rotifers (Brachionus plicatilis, a live-feed organism for finfish larvae) with these FA [369–371]. In addition an increasing body of research into microheterotrophic PUFA production has concentrated on the thraustochytrids [363]. 10.5.1.1 Bacteria In marine food webs, microalgae have long been considered as the only de novo source of EPA [372]. Indeed PUFA were once thought to be absent in bacterial membranes [373] and the production of PUFA by bacteria is often ignored [374]. However, numerous bacterial species of marine origin have now been shown to produce LC-PUFA such as EPA and DHA and some authors correctly pointed to the potential role of prokaryotic PUFA production in marine food webs [25, 347, 375]. Phylogeny The screening of PUFA-producing prokaryote has led to the identification of previously undescribed taxa within the genera Shewanella and Colwellia [376, 377]. Subsequently, the chemotaxonomy of PUFA-producing bacteria was reviewed [30]. Indeed, until recently the taxonomic status of PUFAproducing bacteria received little attention, with most research efforts focused on the description of PUFA production [31]. Figure 22 represents the division-level diversity of the bacterial domain based on 16S rDNA sequences. Several of the divisions (e.g. Proteobacte-

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ria, Actinobacteria) are well represented by cultivated strains. However the majority are poorly represented by cultured organisms. In fact 13 of the 36 divisions are defined by environmental sequences only [378]. The proportion of cultivated versus uncultivated sequences obtained from environmental molecular analyses may be used as an indication of the level of described biodiversity for a particular division. The occurrence of bacteria with the ability to produce PUFA is limited to five well-known marine genera which fall within two distinct domains of bacteria. Although separated by a significant evolutionary distance, the ability to produce EPA is apparent from both groupings. Species from each domain also possess the ability to produce further PUFA products, namely AA or DHA, respectively (Fig. 22) Thus, the ability to produce PUFA exhibits a phylogenetic linkage centred on two distinct lineages, the marine genera of the γ -Proteobacteria (Shewanella, Colwellia, Moritella, Psychromonas, Photobacterium) and more limited species within two genera of the Cytophaga-Flavobacterium-Bacteroides (CFB) grouping (Flexibacter, Psychroflexus). However, it is pertinent to note that not all species within

Fig. 22 Evolutionary distance tree of the bacterial domain [31]. The major PUFAproducing genera, Shewanella, Colwellia and Moritella, are expanded from the Proteobacteria division together with Flexibacter and Psychroflexus from the Cytophagales division. For each genus, biomarker FA and the types of PUFA are listed

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Fig. 23 Schematic of the phylogenetic relationship of the genus Shewanella based on 16S rDNA sequence. Species known to produce polyunsaturated FA (PUFA) and their lines of descent are highlighted in black. Species and lines of descent known not to produce PUFA are highlighted in grey. Arrows indicate sites of divergence where the expression of PUFA synthesis has been lost. Adapted from Russell and Nichols [30]

these genera express the ability to produce PUFA (Figs. 22, 23). Evidence implies that PUFA production is associated with physiological adaptations within marine bacteria. Marine ecology Bacterial PUFA-producing isolates have been found to be particularly prevalent in high-pressure low-temperature deep-sea habitats and permanently cold marine environments [357, 367, 379]. Indeed, the majority of the γ Proteobacteria PUFA producers are characterized as being psychrophilic3 , halophilic and predominantly piezophilic4 or piezotolerant [30, 380]. These physiological traits have influenced the ecological distribution of PUFAproducing bacteria in the marine environment (Table 5). The enrichment of PUFA-producing strains from these environments has led to speculation that PUFA synthesis is an important adaptation for countering the effects of elevated hydrostatic pressure and low temperature on membrane fluidity or phase. In strains which have been analysed, PUFA synthesis undergoes 3

The term psychrophile (psychro = cold, phile = loving) is an operational definition, to describe the temperature-growth relationship of microorganisms [400]. 4 (Piezo = pressure, phile = loving). Same comments as above.

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temperature-dependent and, for deep-sea isolates, pressure-dependent regulation. Typically, as cultivation temperature is decreased, and or pressure increased, PUFA incorporation into membrane phospholipids is enhanced. This modulation is thought to maintain appropriate membrane physical structure. Indeed, in marine bacteria, PUFA are component FA of certain phospholipids which occur in the cell membrane. It is considered that the low melting temperature of these highly unsaturated membrane components, combined with their unique molecular geometry in the membrane, yields a particular advantage at low temperature in balancing the competing homeoviscous and homeophasic forces in the cell membrane [30]. However, if bacteria of the CFB grouping are similarly psychrophilic and halophilic, they lack the ability to grow at high pressure [380]. In addition, for at least one high-pressure-adapted deep-sea bacterium, Photobacterium profundum strain SS9, growth at high pressure and low temperature does not depend upon PUFA synthesis [351]. Hence while PUFA production appears as a phylogenetically linked genotypic strategy for such selective pressures, their presence may not be essential for the growth of bacteria in such environments. While there is not necessarily a phylogenetic relationship between psychrophilic organisms, such relationships may exist through the evolution of adaptive strategies for low-temperature growth. This is supported by empirical observations such as in the genus Shewanella (Fig. 23). Here, there is a good correlation between those species which are cold-adapted and produce PUFA (S. pealeana, S. hanedai, S. benthica, S. gelidimarina, S. frigidimarina), in contrast to those which do not produce PUFA and grow at higher temperatures (mesophiles) (S. putrefaciens, S. alga, S. baltica, S. woodyi, S. oneidensis, S. amazonensis). However, as indicated by Nichols et al. [31], S. frigidimarina and S. pealana are rather psychotolerant than psychrophilic and in addition S. frigidimarina also grows without the presence of salt. Thus these authors point out that the correlation among psychrophily, halophily and PUFA production is therefore not exclusive. They suggest that PUFA production may be a common physiological strategy for coping with the combined constraints of low temperature and marine salinity. In addition to cold environment (sea ice) and deep sea water, PUFAproducing bacteria have also been isolated in the intestinal contents of marine fish and invertebrates [347, 368, 379, 395–397] Moreover, the transfer of bacterially derived FA and specifically PUFA, between marine bacteria and higher trophic levels has been demonstrated [370, 398]. For example, recent studies have demonstrated that PUFA-producing bacteria can be used to enrich rotifers in EPA or DHA [370, 371, 399]. The greatest level of EPA enrichment in rotifers was 5.8% dry weight [370], whereas for DHA it was 0.3% dry weight [371]. These studies provide an important step in the development of novel sources of PUFA for aquaculture [346].

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Table 5 Major bacterial genera responsible for the production of PUFA in the marine environment (adapted from Nichols [25]) Species

Ps

Ha

Pi

PUFA

Environmental source

References

Shewanella S. algae S. amazonensis S. baltica S. benthica S. colwelliana S. frigidimarina S. Gelidimarina S. hadenai S. japonica S. livingstonensis S. oneidensis S. pealeana S. putrefaciens OG1 S. putrefaciens OG3 S. woodyi S. violacea

– – – + – ± + + ± ± – ± – – ± +

– + – + + – + + – – – + – – + +

– – – + – – ± ± – – – – – – – +

– – – + – + + + + nd – + – – – +

Red algae, Japan Water, Amazon river Oil brine, Japan Holourithan intestine Aquaculture, USA Sea ice, Antarctica Sea ice, Antarctica Sediment, Artic Sediments, mussels Sea water, Antarctica Lake sediment, USA Squid gland Butter, UK Butter, UK Sea water, Hawaii Deep-sea

[376] [381] [376] [376] [31] [376] [376] [376] [382] [383] [31] [31] [31] [31] [31] [384]

Colwellia C. demingiae C. hadaliensis C. hornerae C. maris C. psychroerythraea C. psychrotropica C. rossensis

+ + + + + + +

+ + + + + + +

nd + nd nd nd nd Nd

+ nd + + + + +

Sea ice, Antarctica Deep-sea Sea ice, Antarctica Sea water, Japan Flounder eggs Burton lake, Antarctica Sea ice, Antartica

[377] [385] [377] [386] [377] [377] [377]

Moritella M. japonica M. marina M. vavanosii M. viscosus

+ + + +

+ + + +

± ± + ±

+ + + nd

Deep-sea Sea water Deep-sea Fish

[384] [387] [388] [389]

Psychromonas P. antarticus P. kaikoae P. marina

+ + +

+ + +

+ + +

nd + +

Sea-ice, Antarctica Deep-sea Seawater

[390] [391] [392]

– +

– +

nd nd

– +

Burton lake, Antarctica Sea ice, Antarctica

[393] [393]

+

+

+

+

Deep-sea

[394]

Psychroflexus Ps. gondwanense Ps. torquis Photobacterium Ph. profundum

Ps: Psychrophilic, Ha: Halophilic, Pi: Piezophilic

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All theses facts uphold the idea presented by Nichols [25] that the assumption that microalgae provide the bulk of de novo PUFA production for all marine food webs must now be actively reviewed to determine the role and potential importance of PUFA-producing prokaryotes in marine microbial niches such as sea ice, marine animals and abyssal communities. Biotechnology Interest in the production of PUFA from alternative sources for use in aquaculture feeds and human nutraceuticals (see section on heath benefit) has fuelled recent research into the molecular biology of PUFA production in prokaryotes. A key advantage of bacterial PUFA production (as for thraustochytrid PUFA production, see below) is that only a single PUFA is produced, rather than the complex mixture yielded from fish or algal oils [30]. Thus bacterial sources of PUFA remove the expense of preparative purification in the production of high-purity PUFA oils. In addition to their potential use as “cell factories”, bacteria in particular offer the biotechnological opportunity to investigate the genes and enzymes responsible for PUFA production. A variety of bacterial fatty-acid biosynthetic mechanisms exist, which vary with taxonomic identity and class of fatty acid product [401–403]. Some reports have suggested that bacterial (n-3) PUFA production is mediated by undefined desaturases [30, 348, 352]. However, as indicated by Allen et al. [351], sequence studies of bacterial genes required for PUFA biosynthesis have gradually led to a reappraisal of this view (see section on biosynthesis). Initial insight into the genetics of bacterial PUFA synthesis was gained by the transfer of a gene cluster from Shewanella putrefaciens SCRC-2738 into Escherichia coli and a Synechococcus sp. resulted in the successful expression of EPA in these organisms [347]. However, the level of expression achieved was low. The gene cluster used in both cases consisted of a 38 kb fragment containing eight open reading frames with three of these possessing homology with genes that encode for enzymes involved in fatty-acid biosynthesis. Further characterization using these organisms has identified five genes responsible for PUFA biosynthesis, designated ORFs 2, 5, 6, 7 and 8. A subsequent analysis of the predicted amino acid sequences of the products of these genes indicated that they are most related to microbial polyketide synthase (PKS) complexes and fatty acid synthase (FA) enzymes (see section on biosynthesis). In addition to the Shewanella sp. SCRC-2738 sequences, related genes partially responsible for PUFA production have been analysed from the DHA-producing bacterium Moritella marina strain MP-1 (formerly Vibrio marinus) [352] and from a DHA-producing thraustochytrid marine protist belonging to the genus Schizochytrium [329]. Recently, Metz et al. [329] reported biochemical analyses of PUFA production in E. coli strains harbouring Shewanella sp. SCRC-2738 DNA and in the Schizochytrium species. Consistent

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with the examination of enzyme domains, isotopic-labelling studies provided compelling support for a PKS-like pathway of PUFA synthesis in both systems studied [329]. After spending significant effort on understanding the mechanistic microbial production of PUFA, research now has to focus on the regulation of such biosynthesis to maximize the transgenic potential of bacterial LC-PUFA genes. 10.5.1.2 Thraustochytrids Thraustochytrids are microheterotrophs generally considered necessarily marine with a specific requirement for Na+ ions [404] that feed as saprobes or occasionally as parasites [405]. Thraustochytrids have been characterized by the presence of sagenogenetosome, an ectoplasmic net, a cell wall with noncellulosic scales, and a life cycle consisting of vegetative cells, zoosporangium, and biflagellate zoospores [354]. They have a wide geographic distribution in estuarine and marine habitats, with strains isolated from Antarctica [406], the North Sea [407], India [408], Micronesia [409], Japan [410], Australia [359] and Fiji [354]. They have been reported frequently from seawater, sediments, algae, and invertebrates, both as saprotrophs and parasites. Their ubiquitousness and physiological capabilities to utilize a wide variety of habitat suggest that they play a definite role in the marine ecosystem. Originally thought to be primitive fungi, thraustochytrids have more recently been assigned to the subclass Thraustochytridae (Chromista, Heterokonta), aligning them more closely with the heterokont algae (e.g., brown algae and diatoms) [411]. In research on microheterotrophic PUFA production, particular attention has been given to the thraustochytrids since their lipids contains proportionately large quantities of ω3 PUFA and particularly DHA [357]. Bowles et al. [386] have screened 57 thraustochytrids isolates from three different locations. Although a common fatty-acid profile for the thraustochytrid isolates emerged ((n-3) PUFA as a significant component, as previously found [?]), there was considerable difference in the DHA content of the oil. This large variation in DHA proportion can also be extended to biomass and lipid yields depending on the thraustochytrid strains (Table 6). Thus, in some isolates from a cold-temperate environment, DHA can represent almost 50% of the total FA present while those from a sub-tropical environment produce higher levels of biomass, with up to 37 (w/w) % oil but with a lower DHA content [423]. As indicated by Lewis et al. [363], most reports concerning the production of PUFA by thraustochytrids have dealt almost exclusively with DHA production (Table 6), as this compound is often the most abundant PUFA produced by strains of thraustochytrids reported to date. However, it is evident

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that some thraustochytrid strains also produce other PUFA. Thus, Huang et al. [354] have shown that the fatty acid profiles of DHA-producing thraustochytrids could be used to classify them into five separate categories: – – – – –

DHA/DPA (docosapentaenoic acid; C22:5 (n-6)), DHA/DPA/EPA, DHA/EPA, DHA/DPA/EPA/AA DHA/DPA/EPA/AA/DTA (docosatetraenoic acid, C22:4 (n-6)).

Their seven isolates from Japan and Fiji were proved to be new thraustochytrids by their specific insertion sequences in the 18S rRNA genes. The phylogenetic tree constructed by molecular analysis of 18S rRNA genes from those isolates and typical thraustochytrids shows that strains with the same PUFA profile form each monophyletic cluster. These results suggest that the C20–22 PUFA profile may be applicable as an effective characteristic for grouping thraustochytrids. Moreover, Bowles et al. [423] have shown that, among their 57 thraustochytrids, all synthesized the ω6 PUFA arachidonic acid in varying amounts, mainly as a minor component of the PUFA and that EPA was present in the oil produced by all the isolates except two (however EPA content was generally low, varying from 0.2 to [–(%w/w)]0.6 of the dried thraustochytrid cells). So, although about 15 strains of thraustochytrids, including Thraustochytrium aureum, T. roseum, T aggregatum, Schiizochytrium limacinum and S. aggregatum, have been reported to produce significant amounts of DHA (Table 6), there are many potential strains yet to be explored [363, 424]. As indicated by Hammond et al. [425], there are no reports in the literature of direct human consumption of thraustochytrids. This is due to the fact that, prior to the late 1980s, thraustochytrids had never been cultured on a scale larger than a laboratory shake flask. Barclay [422] and Bajpai et al. [414, 415] were the first to successfully cultivate Schizochytrium sp. and Thraustochytrium sp., respectively, in fermenters (two patents have been filed detailing the cultivation of thraustochytrid strains to produce lipids containing EPA and DHA [422, 426]). However, thraustochytrids are primarily consumed by filter-feeding invertebrates in the marine ecosystem, including mussels and clams, and by fish that are consumed directly by humans. Thus, in the last few years, thraustochytrids have been successfully used for commercial production of PUFA-rich products notably in aquaculture applications. This is the case for a Schizochytrium strain, which is the basis for two products marketed for enriching rotifers (Brachionus sp.) and brine shrimp (Artemia sp.) with PUFA, prior to feeding these organisms to cultured finfish larvae ([427]; www.aquafauna.com; www.sandersbshrimp.com). OmegaTech commercialized a product for aquaculture applications (HUFA 2000, a spray-dried form of Schizochytrium sp. dried microalgae), which has been successfully utilized for over seven years as an excellent stable dietary

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source of DHA in shrimp larvaculture and finfish (red seabream, Japanese flounder) culture with no adverse effects. Use of Schizochytrium sp. in these applications has been found to promote larvae survival and growth [428]. Other uses of thraustochytrid oil are being actively explored. Monsanto (www.monsanto.com) is producing Schizochytrium sp.–derived oil under a cooperative technology agreement with OmegaTech (www.omegadha.com). Moreover, dried Schizochytrium microalgae (DRM, OmegaTech) have also been generally recognized as safe (GRAS) for use as a DHA-rich ingredient in broiler chicken and laying-hen feed at levels up to 2.8–4.3%, respectively [429]. Since 1997, DHA-enriched eggs from hens fed a diet containing approximately 1% DRM are now commercially marketed in the United States, Mexico, Germany, Spain, Portugal, the Benelux countries, Italy, Norway, and Israel [430]. These products have entered the market in direct competition with microalgal and fish-oil products. It is possible, however, that thraustochytrids will offer some advantage over other oils as sources of PUFA for aquaculture. Many aquaculture species require proportionally more DHA than EPA in their diet [431]. The PUFA profiles of many thraustochytrids fit this criterion, while most oils from the fish-meal industry contain more EPA than DHA. However it has to be noticed that a recent study [432] has revealed that the replacement of fish oil with a dried product made from a thraustochytrid culture in canola-oil-based diets for Atlantic salmon could affect the disease resistance of fishes. Indeed, if the authors didn’t observe significance differences in final weight, weight gain, feed consumption, feed efficient ration or protein value between the diets, nor in whole-body chemical composition, organ somatic indices or measures of immune function, they have noticed that, following transfer to seawater and two challenges with Vibrio anguillarum, cumulative mortality was significantly lower in fish fed some fish oils than the others. They concluded that their thraustochytrid strain had no detrimental effects on the performance of salmon although, at the current inclusion of 10%, it failed to confer the same effects as fish oil under challenging conditions. In conclusion, as indicated by Lewis et al. [363], thraustochytrids are clearly a new and potentially competitive player in the PUFA market. Considerable work is required before the production of oil from these organisms significantly increases its share of the market for PUFA-rich products. To achieve this aim, the following key stages need to be negotiated. Firstly, the collection, screening, and maintenance of PUFA-producing strains. Several strains with potential for the commercial production of DHA-rich oils have been isolated already. However, if thraustochytrids that produce higher yields, more attractive PUFA profiles, or other less common but sought-after PUFA are isolated and optimized, then demand for these isolates and compounds may well increase. Secondly, the efficiency of PUFA production must be optimized. The types and amounts of PUFA produced by individual strains

2.5

4

5

10

6

Shizochytrium sp. SR21

Shizochytrium sp. SR21

S. Limacium SR21

S. aggregatum ATCC 28209

Thraustochytrium aureum ATCC 34304

Age (d)

25

25

25

28

Temp (◦ C)

Flask 300 rpm

Flask 200 rpm

Flask

Fermenter 300 rpm

Fermenter

Vessel

Light

Dark

–

–

PH. 4

Other

3.8

0.9

38

48

21

Biomass (g/L)

Table 6 Docosahexaenoic acid (DHA) production by thraustochytrids [350]

16.5

–

37

77

50

Lipid (% dw)

49

1.7

33

36

35

(% TFA)

70

30

110

277

224

(mg/g)

270

0.4

4200

13 300

4700

(mg/L)

[414]

[360]

[357]

[413]

[412]

Refs.

100 J.-P. Bergé · G. Barnathan

6

2.5

6

5

12

4

6

T. aureum ATCC 34304

T. aureum ATCC 28211

T. roseum ATCC 28210

T. roseum ATCC 28210

Thraustochytrium sp. ATCC 20892

Thraustochytrium sp. ATCC 20892

Age (d)

T. aureum ATCC 34304

Table 6 (continued)

28

25

25

25

25

25

25

Temp (◦ C)

Flask 120 rpm

Flask 200 rpm

Flask 250 rpm

Flask 250 rpm

Flask 200 rpm

Flask

Flask 300 rpm

Vessel

Light

–

Fed batch

Light

Dark

Light

Light

Other

7.5

2.7

17.1

7.6

0.8

5.7

4.9 20.3 51 104 511 1

Biomass (g/L)

32

7.3

25

18.2

–

8.1

Lipid (% dw)

25

35

49

50

3.7

40

(% TFA)

–+

25

115

87

50

–

(mg/g)

–

68

2100

650

4.0

–

(mg/L)

[419]

[418]

[417]

[416]

[360]

[356]

[415]

Refs.

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of thraustochytrids are susceptible to manipulation by varying culture conditions. Enhancement of PUFA profiles using molecular techniques may be also considered. Different markets will provide demand for strains that produce high levels of PUFA measured either in terms of biomass (i.e., PUFA production w/w cell mass) or volume (i.e., PUFA production w/vol fermentation medium). Thirdly, appropriate conditions for long-term storage of microbial cells and their products must be determined. The form and stability of thraustochytrid biomass and of oils will be major factors in determining the suitability of these products for use as food additives. Finally, oil extraction and refinement technologies must be developed to meet market demands for cost-effective and safe trophic transfer of PUFA to the target consumers. The bottom line for the biotechnological future of thraustochytrid oils will be their competitiveness against other PUFA-rich oils. Nevertheless, oils derived from fish and microalgae generally have a complex fatty acid (total and polyunsaturated) profile, and do not readily lend themselves to the isolation of high-purity (> 98%) FA. Conversely, oils produced by some thraustochytrids have relatively simple fatty-acid profiles and may well be more amenable to cost-effective refinement. DHA is a good illustration of the industrial potential of thraustochytrids. Thus, the largest potential market for microbial oils containing DHA is perceived to be as an additive to infant formulae as an essential fatty acid for brain and retinal development (see section on health benefit); Ratledge [362] considered that the presence of significant quantities of EPA in the thraustochytrid oils so far assessed precluded its use for this purpose. Indeed, eicosapentaenoic acid is considered contraindicatory in breast milk substitutes, but strain selection will easily allow this difficulty to be overcome [423]. 10.5.1.3 Conclusions Growing interest in PUFA applications in various fields coupled with their significance in health and dietary requirements (see section on health benefit) has encouraged “hunting” for more suitable sources of these compounds. The inadequacy of conventional agricultural and animal oils has put attention on developing new microbial technologies. Indeed, microorganisms represent the largest reservoir of undescribed biodiversity, and hence possess the greatest potential for the discovery of new natural products. It is estimated that the Earth currently supports 3–30 million species of organisms. Of these, approximately 1.4 million have been described by science. This includes virtually all the species of birds and mammals (∼ 13 500). In contrast only around 200 000 of the estimated 1.0–1.5 million species of fungi have been characterized (i.e. 13–20%). For the bacteria this percentage is even lower with estimates ranging from only 1–10% of probable species being described in culture [433, 434].

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However, the focus of biotechnology on highly valuable PUFA requires knowledge of how microorganisms control and regulate the fatty-acid biosynthetic machinery in order to obtain specific PUFA in high yield. Elucidation of the signalling systems and mechanisms transmitting the signals from different membranes to the major sites of lipid biosynthetic machinery represents a challenging and potentially rewarding subject for further research [435]. At least, the extensive research and development of PUFA production carried out over the past few years will be aimed at improving the economic competitiveness of microbial lipids compared to plant- and animalderived oils. Nowadays, by using conventional stirred-tank fermenters, economically viable quantities of certain microorganisms that are rich in LC-PUFA can be produced [436]. The chief advantages of such techniques lie in the consistency and purity of the final fatty acid product. Further, unlike the scenario with fish oils, economies of scale have reduced the price of oils derived from organisms raised in fermenters by 10- to 30-fold [437]. 10.5.2 PUFA from fish Fish is a major source of food for mankind, providing a significant amount of the animal protein diet in many countries. Moreover, the consumption of fish has been linked to health benefits (see section on health benefit). Indeed, oils from fish are characterized by a large range of FA from 12–26 carbon atoms and 0–6 double bonds. The bulk of the fatty acid chains is contributed by saturated (15–25%), monoenes (35–60%) and polyenes (25–40%). In contrast with the other fats and oils, fish oils contain large amounts of EPA and DHA, respectively, 14–19% and 5–8%. The proportion of polyunsaturated FA depends on many parameters (see below). Saturated FA include C12 up to C24:0 components, and some branched chains (iso C16, iso C17.) are also found. Among the monoenes, 16:1(n-7), 20:1(n-9) and 22:1(n-11) are present in various amounts, this last component being bioconverted from the corresponding fatty alcohol of copepod wax ester by the fish liver [438]. More than 50 different FA were described in marine fish oil, but eight species frequently represent more than 80% of the total amount (Table 7). In fish tissues, the composition of FA (mainly of triacylglycerols and to a lesser extend of phospholipids), is determined by diet composition and lipid metabolism [439, 440]. Fish have the ability to synthesize de novo the saturated and monounsaturated FA and also to selectively absorb and metabolize dietary FA including LC-PUFA [440, 441] in order to obtain an optimal fatty acid composition [442]. This optimal composition seems to be a characteristic for each species and even each strain [443, 444]. Moreover, the PUFA conversion capacity in fish varies among species and even races [439]. Thus, freshwater fish are generally able to elongate and desaturate α-linolenic acid

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Table 7

14:0 16:0 16:1(n-7) 18:0 18:1(n-9) 20:1(n-9) 20:5(n-3) 22:1(n-11) 22:6(n-3)

Menhaden

Herring

7–12 15–26 9–16 2–4 8–14 – 11–16 <1 5–14

5–8 10–19 6–12 1–2 9–25 7–20 4–15 7–30 2–8

(18:3(n-3)) to EPA and DHA, whereas marine fish, which lack or have a very low activity of ∆5-desaturase, cannot and require LC-PUFA such as EPA and DHA in the diet [440]. In addition to food accessibility and lipid metabolism some environmental parameters also notably influence the proportion of PUFA [445]. Indeed, the colder the water, the higher the amount of these components. An inherent property of cells in poikilothermic animals is their capacity to adjust the physicochemical characteristics of their membranes to the prevailing temperatures. This phenomenon, known as homeoviscous adaptation of membrane fluidity, has been reported in poikilothermic fish [446]. In fish, during adaptation to reduced temperatures, unsaturation of the constituent FA increases, with the polar head group as well as the molecular species composition of membrane phospholipids being reorganized [202]. Evidence suggests that the fatty acid distribution is very individual from species to species and depends on many factors like season, temperature, fishing ground, fish species, age, gender or nutritional habits [447–452]. The FA distribution in triacylglycerols is stereospecific [www.cyberlipid. org]. Indeed, results indicate that the stereospecific location of the PUFA is in position 2 (also 3 for EPA in cod) in fish but in position 3 for mammals (same in seal, whale and polar bear). This raises the question of the availability of these FA from food expecting a benefic effect on diverse human functions (see section on health benefit). Familiar fish species used in the production of fish oil include among others, anchovies, capelin, Atlantic cod, Atlantic herring, Atlantic mackerel, Atlantic menhaden, salmonids, sardines (see section on market and sources). Of the world’s fish oil production, 90% is produced from fatty fish where lipids are localized mainly under the skin, around the intestines or in the white muscle. In such fishes, the oil content varies (see above) but it can reach 21% (herring) and 18% (sardines). Such oils are still the least-expensive natural source of preformed long-chain PUFA, and several industries (e.g., Ocean

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Nutrition, Halifax, N.S., Canada, and Pronova Biocare, Sandefjord, Norway) now specialize in their production and purification through cold pressing, further concentration by winterization (i.e., chilling), and other technologies [431]. There are, however, potential problems associated with fish oils as a source of PUFA such as: taste, odor, stability problems as well as the presence of coextracted contaminants. Some could be at least partially solved for example by microencapsulation [453] and deodorization [454]. Nevertheless, the crucial problem of those oils is their sustainability due to the worldwide decline of fish stocks (see section on market). A better use of raw material but as well of by-catch and by-products from fisheries may be one solution, another is to look for other sources (see above) including non- or little-exploited fish species. Lipid content has been studied for a long time mainly in the liver and edible parts of fish such as muscles. Several investigations have evidenced that other parts i.e. liver and skin, often discarded for several reasons when fish are prepared for consumption, may have possible nutritional and therapeutic value mainly due to their EPA and DHA content [445, 455]. Indeed valuable fish oils can be obtained from trash fish and/or fish scraps or cannery wastes, which are often called “gurry” from filleting and canning operations. Fish scraps normally consist of the head, skeleton, and adhering proteinaceous tissues. Surprisingly, it can also be observed that often no detailed chemical studies of the lipid and fatty acid content of certain fish are available, even though the fish most often consumed or commercially exploited, such as for example the species Sardinella and Cephalopholis in Senegal waters [455]. Capelin, squid liver, krill (Euphausia superba) could constitute sustainable new sources [437]. 10.6 Current utilization of marine oils and lipids 10.6.1 Market 10.6.1.1 Production From a world production of oil and fat of about 20 million tons in 1939, about 77 million tons were produced in 1989 where 74% were of vegetal origin (soybean 19% > palm 13% > rapeseed 10.3% > sunflower 9.6%). In 2003–2004 the global production of fats and oils is expected to be 128.5 million tons with 82% of vegetal origin. The world average consumption of oils and fats in 2003 is about 20 kg per capita [www.cyberlipid.org].

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Of the estimated 89 million tons of fish produced in 2000 in the world, excluding China, nearly 71 percent (63 million tons) was used for direct human consumption. The remainder (about 29 percent) was utilized for various nonfood products, mostly for reduction to meal and oil (the state of world fisheries and aquaculture, 2002 FAO). Indeed, nowadays, a third of the world’s catch from the seas is going into manufacturing fish meal and fish oil. Thus, the world production of marine oils represent approximately 1% of the commodity world fats and oils production (Table 8). Remark: the International Fishmeal and Fish Oil Organisation (IFFO) is the international nongovernmental trade organization representing fish meal and oil producers worldwide. It has more than 200 member companies in 38 countries. Two-thirds of the world’s production of fish meal and fish oil are members of the IFFO and 95% of the exports of fish meal and oil are also part of the IFFO. Chile, Peru, Scandinavia, USA and Japan are the main suppliers of fish oil (Fig. 24). The average world production between 1991 and 2001 was about 1.25 million tons of fish oil produced annually. Important fluctuations in production can be observed, they were due to the El Niño phenomenon mostly in Chile and Peru. In 1998, which was the big El Niño year, the production of these two countries only reached 210 000 tons while the average over the last five years was 520 000 tons (the total catch volume by Peruvian fisheries was reduced to 3696 million tons, only 45% of the 2002 catch of 8238 million tons). However, due to better prediction of such climatic occurrence, the governments concerned are increasingly proactive in anticipating and taking precautionary measures for fishing. Thus, precautionary approach to fisheries management has been maintained (in Peru notably) to safeguard the viability and prevent depletion of stocks through overfishing. Careful management of the fishery and a return to normal environmental conditions allowed stocks to recover in 1999 and 2000.

Table 8 Production (million tonnes) for 17 commodity oils in the four-year period 1998/99 to 2001/02 [456]

Total production Soybean Palm Rapeseed Sunflower Other vegetal oils Fish Other animal fats and oils

98/99

99/00

00/01

01/02

107.6 24.60 19.40 12.70 9.30 19.70 0.86 21.04

113.5 25.30 21.30 14.50 9.50 20.50 1.38 21.02

117.3 27.10 23.70 13.90 8.70 21.70 1.42 20.78

119.7 29.40 24.30 13.40 7.50 22.40 1.12 21.58

Fatty acids from marine I

107

Fig. 24 World fish body oil production – major producers (IFFO)

10.6.1.2 Exportation The major exporters are mainly the same countries, with the noticeable exception of Japan that is rather now a net importer (Fig. 25). Over the past

Fig. 25 World marine oils and fats exports by major exporters (IFFO)

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J.-P. Bergé · G. Barnathan

decade fish oil exports by Peru (the main exporting country) have expanded by almost twelve times (US$ 91.1 million in 2001). However, Peru’s exportation is variable; it can be enormous (up to 500 000 tons in 2000) or very small, notably during El Niño periods. The second main fish oil exporter is now the USA (US$ 41.7 million in 2001). 10.6.1.3 Consumption Consumption of fish oils by countries is indicated in Fig. 26. Most of the fish oil goes into salmonid feed in Norway, Chile, Canada and various European countries, which is the reason for the predominance of these countries in terms of consumption. Remark: fish oil is included in aquaculture feeds as a source of both dietary energy and PUFA. Considerable research is occurring worldwide in an effort to find alternatives to fishmeal and fish oil in aquaculture feeds. However, this research is tempered by the obligate dietary requirement of many marine finfish species for long-chain PUFA (LC-PUFA: e.g., EPA and DHA). Aquaculture has been the world’s fastest-growing food production over a decade [438]. The world aquaculture production has at least multiplied by a factor of two in the last ten years: 24 457 421 tons live weight in 1993 and 48 413 636 tons live weight in 2001 (Eurostat and FAO sources). The International Fishmeal and Oil Manufacturers Association [www.iffo.com] estimates

Fig. 26 World marine oil consumption and stocks – major consumers (IFFO)

Fatty acids from marine I

109

that inclusion of fish oil in aquaculture feeds will rise from 380 000 tons in 1994 to 582 000 tons in 2001 and 1 133 000 tons in 2010 (Table 9). With aquafeed demand at about 1 million tons of fish oil in 2010, depending on the production of fish oil it could be around 80% or even close to 100%. This could well result in a worldwide undersupply of fish oil, leading to increased demand for fish oil alternatives. Moreover, this lack of fish oil will have an impact on aquafeed composition. It seems likely that cheap, plant- or animal-derived oils, which often contain low levels of LC-PUFA, will be used increasingly as alternative sources of energy in some aquaculture feeds. If such substitution does occur, sufficient LC-PUFA to meet the dietary requirements of cultured aquaculture species may be required from other sources. Typically, many cultured marine species require around 1% to 2 wt/wt % LCPUFA in their diets [457, 458]. Pike and Barlow [459] estimated that marine aquaculture finfish species will require about 2 × 106 tons of feed in 2010. These figures point to a potential demand, for these species alone, for at least 10 000 tons of LC-PUFA per annum (Table 10). In addition to aquafeed, the current and potential world market for fish oil products spans a number of sectors from unprocessed, oil-rich biomass for animal feeds, to high-quality food-grade oils for use as food additives and nutraceuticals, and to very-high-purity oils and even individual FA for use in the pharmaceutical industry (Table 9). As indicated by Lewis et al. [363], the imprecise boundaries surrounding the nutraceutical market make estimating the size of this market sector more difficult. Sales of marine supplement oils were in the order of $55 million in the United States in 1996 [460], and represented 20% of sales from health food retail outlets. In the United Kingdom, fish oils account for approximately 29% (U.S. $140 million) of the total annual market for nutraceuticals [461]. Moreover, there is an increasing trend for infant formula manufacturers to include PUFA-rich oils in their products. Typical inclusion levels of PUFA-rich oils are designed to achieve a final DHA concentration in dry infant formula of 0.1% to 0.2 wt/wt %. Indeed, the Western European market for infant formula increased from 81 500 tons in 1988 to 103 933 tons in 1994. Extrapolating these figures suggests a potential annual demand in the European infant formula

Table 9 Fish oil use prediction based on an annual world production of 1.25 million tonnes for the period 2002–2010 [461]

Edible Industrial Aquafeed Pharmaceutical

1990

2002

2010

76% 8% 16% –

30% 12% 56% 2%

14% 5% 79% 2%

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J.-P. Bergé · G. Barnathan

Table 10 Predicted use of fish oil in fish feed (data from IFFO) % of fish oil inclusion in feed produced Thousand tons of fish oil 2000 2010 2000 2010

Carp Catfish Tilapia Milkfish Shrimp Eel Marine fish1 Trout Marine fish2 Salmon TOTAL 1 2

– 1.0 1.0 2.0 2.0 5.0 10.0 15.0 20.0 25.0

0.5 – 0.5 2.0 3.0 8.0 12.0 15.0 15.0 20.0

– 5 8 6 30 17 23 95 226 307

103 – 9 11 73 23 156 121 335 379

716

1209

Flat fish including flounder, turbot, halibut, sole and cod, hake Bass, bream, yellowtail, grouper, jacks, mullets

market for up to 100 to 200 tons of DHA. Several food and beverage products enriched with DHA or other PUFA are already on the market. Mukherjee [461] reported the availability of products such as enriched spreads, breads, eggs, and soft drinks in Europe and Japan. Bread enriched with refined tuna oil as a source of LC-PUFA is achieving substantial market penetration in Australia. As awareness by both consumers and regulators of the importance of adequate levels of PUFA in our diet increases, it can be assumed that demand for a greater range of PUFA-enriched products will increase [363]. 10.6.1.4 Prices The biggest use of fish oils is by the aquaculture industry, where it is necessary to have an oil rich in the long-chain polyunsaturated FA characteristic of fish oils. For this purpose, therefore, fish oil cannot be adequately replaced by vegetable oils. To meet this demand there has been a reduction in stocks and an increase in price. In 2000 and 2001 the average monthly price for crude fish oil ranged from US$ 235–325/ton and $ 323–598/ton. In January 2002 it was $613/ton. [Oil World, www.oilworld.org]. In August 2002, crude fish oil prices peaked (about 650 US$ per ton) and have started to decline ever since. In November 2002, finally soybean oil prices managed to overtake those of crude fish oil due to the shorter supply than initially forecast, which make the latter competitive once more on the hardening market. For 2002, the average price of crude fish oil from any origin was about US $587/ton; in 2003 it was

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about US$ 562/ton (data from Oilworld). Peru’s enormous fish oil production capacity sets this product’s international prices. 10.6.2 Common resources Fish oil is a by-product of industrial fishing and the fish meal industry. Fish oils are produced almost exclusively from small, bony species of pelagic fish (living in the surface waters or middle depths of the sea), for which there is little or no demand for human consumption [Fishmeal Information Network, www.fin.org.uk]: South America (three species) In Peru, anchovy is by far the most important species for fishmeal and fish oil production, with sardine largely making up the difference. The Chilean fishmeal industry uses anchovy, sardine and jack mackerel. Europe (seven species) Seven key species are used to produce fishmeal and fish oil in Europe. These can be divided into three groups: a) No use for human consumption (inedible feed-grade fish – sandeel, capelin, Norway pout). b) Potential use for human consumption but mainly used for fishmeal because of limited outlets for human consumption (blue whiting, sprat). c) Primary use is human consumption but surplus may be used for fishmeal (herring, horse mackerel). Fishmeal production also provides a major outlet to recycle trimmings from the food-fish processing sector which would otherwise be dumped at extra cost to the environment and the consumer. In the EU, Spain, France, Germany, Ireland and the UK produce fishmeal and fish oil primarily from trimmings. Global capture fisheries, i.e. catches of wild fish as distinct from farmed fish, are valuable and finite resources which, although renewable, are highly vulnerable. Moreover, overfishing has caused the collapse or near collapse of some valuable fisheries. Overexploiting one fish species can affect other species, not least birds and mammals, in the marine ecosystem. This situation has generated understandable and justifiable pressure for environmentalists to reduce fishing effort and catches further by introducing tighter regulatory measures [438].

References 1. Sargent JR (1995) Origins and functions of egg lipids: nutritional implications. In: Bromage NR, Roberts RJ (eds.), Broodstock Management and Egg and Larval Quality. Blackwell Science, Oxford, p 353

112

J.-P. Bergé · G. Barnathan

2. Sargent JR, Bell MV, Bell JG, Henderson RJ, Tocher DR (1995) Origins and functions of n-3 polyunsaturated fatty acids in marine organisms. In: Cevc G, Paltauf F (eds.), Phospholipids: Characterization, Metabolism and Novel Biological Applications. AOCS, Champaign, Illinois, p 248 3. Pazos AJ, Roman G, Acosta CP, Sanchez JL, Abad M (1997) Comp Biochem Physiol 117B:393 4. Ackman RG (ed.) (1989) Marine Biogenic Lipids Fats and Oils. Vol I II. CRC, Boca Raton, Florida 5. Rezanka T (1989) Progr Lipid Res 28:147 6. Christie WW (ed) (2003) Lipid Analysis, Isolation, Separation, Identification and Structural Analysis of Lipids. The Oily Press, PJ Barnes, Brigwater, UK 7. Hagen W, Auel H (2001) Zoology 104:313 8. Dalsgaard J, St John M, Kattner G, Müller-Navarra D, Hagen W (2003) Adv Mar Biol 46:227 9. Fattorusso E, Mangoni A (1997) Marine Glycolipids. In: Hertz W, Kirby GW, Moore RE, Steiglich W, Tamm W (eds), Progress in the Chemistry of Organic Natural Products. Springer, Wien, p 215 10. Costantino V, Fattorusso E, Mangoni A, Di Rosa M, Ianaro A (1997) J Am Chem Soc 119:12 465 11. Costantino V, Fattorusso E, Imperatore C, Mangoni A (2003) Eur J Org Chem 1433 12. Crul M, Mathot RAA, Giaccone G, Punt CJA, Rosing H, Hillebrand MJX, Ando Y, Nishi N, Tanaka H, Schellens J, Beijnen JH (2002) Cancer Chemotherap Pharmacol 49:287 13. Ackman RG (2002) Anal Chim Acta 465:175 14. Napolitano GE, Pollero RJ, Gayoso AN, MacDonald BA, Thompson RJ (1997) Biochem Syst Ecol 25:739 15. Zhukova NV, Titlyanov EA (2003) Phytochemistry 62:191 16. Hooper JNA, Van Soest RWM (2002) Systema Porifera: A Guide to the Classification of Sponges, Kluwer Academic/Plenum, New York 17. Gurr MI, Hardwood JL (1991) Lipid Biochemistry: An Introduction. Chapman Hall, London, UK 18. Cook HW (1996) Fatty acid desaturation and chain elongation in eukaryotes. In: Biochemistry of Lipids, Lipoproteins and Membranes. Vance DE, Vance JE (eds), Elsevier Scientific, Amsterdam, p 129 19. Sargent JR, Henderson RJ (1986) Lipids. In: The biological chemistry of marine copepods, Vol 1, EDS Corner and SCM O’Hara (eds.), Clarendon, Oxford, UK, p 59 20. Kattner G, Hagen W (1995) ICES J Mar Sci 52:329 21. Sprecher H (1992) Omega-3 News 7:1 22. Sprecher H (2000) Biochim Biophys Acta 1486:219 23. Williard DE, Nwankwo JO, Kaduce TL, Harmon SD, Irons M, Moser HW, Raymond GV, Spector AA (2001) J Lipid Res 42:501 24. Henderson RJ (1999) The production of n-3 polyunsaturated faty acids in marine organisms. In: Lipid Technology, Barnes PJ, UK, p 5 25. Nichols DS (2003) FEMS Microbiol Lett 219:1 26. Nadjek M, Debobbis D, Miokovic D, Ivancic I (2002) J Plankton Res 24:494 27. Harvey HR, Macko SA (1997) Organic Geochem 26:531 28. Budge SM, Parrish CC (1998) Organic Geochem 29:1547 29. Budge SM, Parrish CC, McKenzie CH (2001) Mar Chem 73:285 30. Russel NJ, Nichols DS (1999) Microbiology 145:767 31. Nichols DS, McMeekin TA (2002) J Microbiol Meth 48:161

Fatty acids from marine I

113

32. Pond DW, Allen CE, Bell MV, Van Dover CL, Fallik AE, Dixon DR, Sargent JR (2002) Mar Ecol Progr Ser 225:219 33. Yu R, Yamada A, Watanabe K, Yazawa K, Takeyama H, Matsunaga T, Kuranr R (2000) Lipids 35:1061 34. Hannun YA, Obeid LM, Dbaibo GS (1996) Handbook of Lipid Research, Vol 8, p 177 35. Wha Sona B, Chul Kima J, Dae Choib H (2001) Lipids 36:427 36. Dembitsky VM, Srebnik M (2002) Progr Lipid Res 41:315 37. Wu M, Milligan KE, Gerwick WH (1997) Tetrahedron 53:15 983 38. Kan Y, Sakamoto B, Fujita T, Nagai H (1998) J Nat Prod 61:152 39. Mesguiche V,Valls R, Piovetti L, Peiffer G (1999) Tetrahedron Lett 40:7473 40. Kan Y, Sakamoto B, Fujita T, Nagai H (2000) J Nat Prod 63:1599 41. Falk-Petersen S, Sargent JR, Henderson J, Hegseth EN, Hop H, Okolodkov YB (1998) Polar Biol 20:41 42. Henderson RJ, Hegesth EN, Park MT (1998) Polar Biol 20:48 43. Volkman JK, Barett SM, Blackburn SI, Mansour MP, Sikes EL, Gelin F (1998) Organic Geochem 29:1163 44. Reuss N, Poulsen LK (2002) Mar Biol 141:423 45. Viso AC, Marty JC (1993) Phytochemistry 34:1521 46. Viron C, Saunois A, André P, Perly B, Lafosse M (2000) Anal Chim Acta 409:257 47. Mostaert AS, Karsten U, Hara Y, Watanabe MM (1998) Phycol Res 46:213 48. Cho ES, Rhodes LL, Kim HK (1999) J Fish Sci Tech 2:58–65 49. Marshall JA, Nichols PD, Hallegraeff GM (2002) J Applied Phycol 14:255 50. Khozin-Goldberg I, Zheng Yu H, Adlerstein D, Didi-Cohen S, Heimer YM, Cohen Z (2000) Lipids 35:881 51. Budge SM, Parrish CC (1999) Phytochemistry 52:561 52. Bergé JP, Gouygou JP, Dubacq JP, Durand P (1995) Phytochemistry 39:1017 53. Xu XQ, Beardall J, Hallam ND (1998) Phytochemistry 49:1249 54. Bell MV, Dick JR, Pond DW (1997) Phytochemistry 45:303 55. Arzul G, Gentien P, Bodennec G, Toularastel F, Youenou A, Crassous MP (1998) Potential toxicity of microalgal polyunsaturated fatty acids. In: Baudimant G, Guezennec JH, Roy P, Samain JF (eds.), Marine Lipids, IFREMER, Plouzané, France, p 53 56. Bodennec G, Gentien P, Parrish CC, Crassous MP (1998) Lipid class and fatty acid compositions of toxic Gymnodinium and Heterosigma strains: haemolytic and signature compounds. In: Baudimant G, Guezennec JH, Roy P, Samain JF (eds.), Marine Lipids, IFREMER, Plouzané, France, p 66 57. Mansour MP, Volkman JK, Holdsworth DG, Jackson AE, Blackburn SI (1999a) Phytochemistry 50:541 58. Mansour MP, Volkman JK, Jackson AE, Blackburn SI (1999b) J Phycol 35:710 59. Rezanka T (1990) J Chromatogr 513:344 60. Van Pelt CK, Huang MC, Tschanz CL, Brenna JT (1999) J Lipid Res 40:1501 61. Tanaka I, Matsuoka S, Murata M, Tachibana K (1998) J Nat Prod 61:685 62. Li X, Fan L, Lou Q (2002) Phytochemistry 59:157 63. Fleurence J, Gutbier G, Mabeau S, Leray C (1994) J Applied Phycol 6:527 64. Khotimchenko SV, Vaskovsky VE, Titlyanova TV (2002) Botanica Marina 45:17 65. Graeve M, Kattner G, Wiencke C, Karsten U (2002) Mar Ecol Progr Ser 231:67 66. Sajiki J, Kakimi H (1998) J Chromatogr 795A:227 67. Kornprobst JM, Barnathan G (1998) Recent Res Dev Lipid Res 2:371 68. Carballeira NM (2002) Progr Lipd Res 41:433 69. Barnathan G, Bourgougnon N, Kornprobst JM (1998) Phytochemistry 47:761

114

J.-P. Bergé · G. Barnathan

70. Smith SL, Schnack-Schiel SB (1990) Polar zooplankton. In: Smith WO (ed), Polar Oceanography, Part B: Chemistry, Biology and Geology, Academic, San Diego, California, p 527 71. Albers CS, Kattner G, Hagen W (1996) Mar Chem 55:347 72. Atkinson A, Snyder R (1997) Mar Ecol Progr 160:63 73. Mayzaud P, Albessard E, Cuzin-Roudy J (1998) Mar Ecol Progr Ser 173:19 74. Pond DW, Sargent JR (1998) J Plankton Res 20:169 75. Kattner G, Hagen W, Graeve M, Albers C (1998) Mar Chem 61:219 76. Kattner G, Hagen W (1998) Mar Ecol Progr Ser 170:203 77. Falk-Petersen S, Sargent JR, Lonne OJ, Timofeev SF (1999) Polar Biol 21:37 78. Falk-Petersen S, Hagen W, Kattner G, Clarke A, Sargent J (2000) Can J Fish Aquat Sci 57:178 79. Falk-Petersen S, Dahl TM, Scott CL, Sargent JR, Gulliksen B, Kwasniewski S, Hop H, Millar RM (2002) Mar Ecol Progr Ser 187:187 80. Scott CL, Falk-Petersen S, Dahl TM, Sargent JR, Hop H, Lonne AJ, Poltermann M (1999) Polar Biol 21:65 81. Scott CL, Kwasniewski S, Falk-Petersen S, Millar RM, Sargent JR, (2000) Polar Biol 23:510 82. Jones PJH, MacDougall DE, Ntanios P,Vanstone CA, Mayzaud P, Albessard E, CuzinRoudy J (1998) Mar Biol Progr Ser 173:149 83. Cripps GC, Watkins JL, Hill HJ, Atkinson A (1999) Mar Ecol Progr Ser 181:177 84. Cho KW, Shin J, Jung K (1999) Ocean Res 21:109 85. Nelson MM, Phleger CF, Mooney BD, Nichols PD (2000) Lipids 35:551 86. Nelson MM, Mooney B, Nichols PD, Phleger CF (2001) Mar Chem 73:53 87. Phleger CF, Nelson MM, Mooney B, Nichols PD (2000) Polar Biol 23:329 88. Phleger CF, Nichols PD, Virtue P (1998) Comp Biochem Physiol 120B:311 89. Phleger CF, Nelson MM, Mooney B, Nichols PD (1999) Comp Biochem Physiol 124B:295 90. Swadling KM, Nichols PD, Gibson JAE, Ritz DA (2000) Mar Ecol Prog Ser 208:171 91. Litchfield C, Morales RW (1976) Are Demospongiae membranes unique in living organisms? In: Harrison FW, Cowden RR (eds.), Aspects of Sponge Biology, Academic, New York, p 183 92. Bergquist P (ed) (1978) Sponges. Hutchinson University Library, London 93. Djerassi C, Lam WK (1991) Acc Chem Res 24:69 94. Christie WW, Brechany EY, Marekov IN, Stefanov KL, Andreev SN (1994) Comp Biochem Physiol 109B:245 95. Carballeira NM, Emiliano A, Hernandez-Alonso N, Gonzalez FA (1998) J Nat Prod 61:1543 96. Barnathan G, Kornprobst JM (1998) Fatty acids from marine organisms: recent research developments. In: Baudimant G, Guezennec J, Roy P, Samain JF (eds), Marine Lipids. IFREMER, Brest, France, p 35 97. Barnathan G, Kornprobst JM (1998) Recent Res Devel Lipids Res 2:235 98. Carballeira NM, Shalabi F (1993) J Nat Prod 56:739 99. Barnathan G, Kornprobst JM, Doumenq P, Mirallès J (1995) Lipids 31:193 100. Carballeira NM (1997) Recent Res Develop Lipid Res 1:9 101. Carballeira NM, Alicea J (2001) Lipids 36:83 102. Barnathan G, Genin E, Velosaotsy NE, Kornprobst JM, Al-Lihaibi S, Al-Sofyani, Nongonierma R (2003) Comp Biochem Physiol 135B:297 103. Makarieva TN, Santalova EA, Gorshkova IA, Dmitrenok AS, Guzii AG, Gorbach VI, Svetashev VI, Stonik VA (2002) Lipids 37:75

Fatty acids from marine I 104. 105. 106. 107. 108. 109. 110. 111. 112. 113. 114. 115. 116. 117. 118. 119. 120. 121. 122. 123. 124. 125. 126. 127.

128. 129. 130. 131. 132. 133. 134. 135. 136. 137. 138.

115

Rodkina SA, Latyshev NA, Imbs AB (2003) Russ J Bioorg Chem 29:382 Meyer M, Guyot M (2002) Lipids 37:1109 Nemoto T, Yoshino G, Ojika M, Sakagami Y (1997) Tetrahedron 53:166 699 Li Y, Ishibashi M, Sasaki T, Kabayashi J (1995) J Chem Res (S):126 Li Y, Ishibashi M, Sasaki T, Kabayashi J (1995) J Chem Res (M):901 Carballeira NM, Medina JR (1994) J Nat Prod 57:1688 Carballeira NM, Reyes M (1995) J Nat Prod 58:1689 Carballeira NM, Reyes ED, Sostre A, Rodriguez AD, Rodriguez JL, Gonzales FA (1997) J Nat Prod 60:502 Kawakami A, Miyamoto T, Higuchi R, Uchiumi T, Kuwano M, Van Soest RWM (2001) Tetrahedron Lett 42:3335 Carballeira NM, Pagan M (2000) J Nat Prod 63:666 Nechev J, Christie WW, Robaina R, de Diego F, Popov S, Stefanov K (2002) Eur J Lipid Sci Technol 104:800 Nechev J, Christie WW, Robaina R, de Diego FM, Ivanova A, Popov S, Stefanov K (2002) Hydrobiologia 489:91 Nechev J, Christie WW, Robaina R, de Diego F, Popov S, Stefanov K (2004) Comp Biochem Physiol 137A: 365 Carballeira NM, Pagan M (2001) J Nat Prod 64:620 Thiel V, Jenisch A, Worheide G, Lowenberg A, Reitner J, Michaelis W (1999) Org Geochem 30:1 Kornprobst JM, Barnathan G (2000) Recent Res Progr Lipid Res 2:371 Ayanoglu E, Kornprobst JM, Aboud-Bichara A, Djerassi C (1983) Tetrahedron Lett 24:1111 Ayanoglu E, Popov S, Kornprobst JM, Aboud-Bichara A, Djerassi C (1983) Lipids 18:830 Carballeira NM, Alicea J (2002) Lipids 37:305 Carballeira NM, Colon R (1999) Tetrahedron Asymmetry 10:3785 Soderquist JA, Rosado I, Marrero Y (1998) Tetrahedron Lett 39:3115 Quinn RJ, Tucker DJ (1985) Tetrahedron Lett 26:1671 Hirsch SC, Carmely S, Kashman Y (1987) Tetrahedron 43:3257 Van Soest RWM, Fusetani N, Andersen RJ (1997) Straight-chain acetylenes as chemotaxonomic markers of the marine Haplosclerida. In: Watanabe Y, Fusetani N (eds), Sponge Sciences–Multidisciplinary Perspectives, Springer-Verlag, Berlin Heidelberg New York, p 3 Pham NB, Butler MS, Hooper JNA, Moni RW, Quinn RJ (1999) J Nat Prod 62:1439 Matsunaga S, Okada Y, Fusetani N, Van Soest RWM (2000) J Nat Prod 63:690 Lee HS, Rho JR, Sim CJ, Shin J (2003) J Nat Prod 66:566 Fujita M, Nakao Y, Matsunaga S, Van Soest RWM, Itoh Y, Seiki M, Fusetani N (2003) J Nat Prod 66:569 Zhao QC, Lee SY, Hong JK, Lee CO, Im KS, Lee DS, Sim CJ, Jung JH (2003) J Nat Prod 66:408 Hattori T, Adachi K, Shizuri Y (1998) J Nat Prod 61:823 Venkateswarlu Y, Srinivasa Reddy N, Ramesh P, Rama Rao M, Siva Ram T (1998) Indian J Chem 37B:1264 Lo JM, Wang WL, Chiang YM, Chen CM (2001) J Chin Chem Soc 48:821 Natori T, Morita M, Akimoto K, Koezuka Y (1994) Tetrahedron 50:2771 Li HY, Matsunaga S, Fusetani N (1995) Tetrahedron 51:2273 Koezuka Y, Motoki K, Sakai T, Natori T (1999) Recent Res Dev Cancer 1:341

116

J.-P. Bergé · G. Barnathan

139. Costantino V, Fattorusso E, Mangoni A, Di Rosa M, Ianaro M (1997) J Am Chem Soc 119:12 465 140. Costantino V, Fattorusso E, Mangoni A (2000) Tetrahedron 56:5953 141. Costantino V, Fattorusso E, Mangoni A (2001) Glycolipids with immunomodulating activity from marine sponges. In: Tringali C (ed.), Bioactive Compounds from Natural Sources. Taylor Francis, London, New York, Chap 14:557 142. Meguro S, Namikoshi M, Kobayashi H (2002) J Antibiotics 55:256 143. Hayakawa Y, Rovero S, Forni G, Smyth MJ (2003) Proc Nat Acad Sci 100:9464 144. Kobayashi E, Motoki K, Uchida T, Fukushima H, Koezuka Y (1998) Oncol Res 7:529 145. Morita M, Motoki K, Natori T, Sakai T, Sawa E, Yamaji K, Koezuka K, Kobayashi E, Fukushima H (1995) J Med Chem 38:2176 146. Costantino V, Fattorusso E, Imperatore C, Mangoni A (2002) J Nat Prod 65:883 147. Vysotskii MV, Svetashev VI (1991) Biochim Biophys Acta 1083:161 148. Vysotskii MV, Svetashev VI (1998) Comp Biochem Physiol 119B:73 149. Mirallès J, Barnathan G, Galonnier R, Sall T, Samb A, Gaydou EM, Kornprobst JM (1995) Lipids 30:459 150. Carballeira NM, Sostre A, Rodriguez AD (1996) Comp Biochem Physiol 113B:781 151. Carballeira NM, Sostre A, Rodriguez AD (1997) Comp Biochem Physiol 118B:257 152. Carballeira NM, Emiliano A, Rodrıguez J, Reyes E (1992) Lipids 27:681 153. Barnathan G, Doumenq P, Mirallès J, Boury-Esnault N, Kornprobst JM (1993) J Nat Prod 56:2104 154. Carballeira NM, Miranda C, Rodrıguez AD (2002) Comp Biochem Physiol 131B:83 155. Urvois PA, Barnathan G, Biard JF, Débitus C, Verbist JF (1998) Fatty acid composition of the New Caledonian gorgonian Rumphella aggregata: identification of 9-methyl-6,9-heptadecadienoic acid. In: Marine Lipids, Baudimant G, Guezennec J, Roy P, Samain JF (eds.), IFREMER, Brest, p 44 156. Mancini I, Guerriero A, Bakken T, Zibrowius G, Pietra F (1999) Helv Chim Acta 82:677 157. Henderson RJ, Burkow IC, Buzzi M, Bayer A (1998) Biochim Biophys Acta 1392:309 158. Henderson RJ (2000) J Mar Biol Ass UK 80:311 159. Yamashiro H, Oku H, Higa H, Chinen I, Sakai K (1999) Comp Biochem Physiol 122B:397 160. Jeong TS, Ahn JA, Kim YK, Bok SH, Kwon BM, (1999) Bioorganic Medicinal Chem Lett 7:1481 161. Ota T, Chihara Y, Itabashi Y, Tagaki T (1994) Fisheries Sci 60:171 162. McKenzie JD, Black KD, Kelly MS, Newton LC, Handley LL, Scrimpgeour CM, Raven JA, Henderson RJ (2000) J Mar Biol Ass UK 80:311 163. Kawasaki KI, Nabeshima YI, Ishihara K, Kaneniwa M, Ooizumi T (2000) Fisheries Sci 66:614 164. Ishihara K, Murata M, Kaneniwa M (1998) Lipids 33:1107 165. Sato D, Ando Y, Tsujimoto R, Kawasaki K (2001) Lipids 36:1371 166. Inagaki M, Isobe R, Kawano Y, Miyamoto T, Komori T, Higuchi R (1998) Eur J Org Chem 129 167. Yamada K, Matsubara R, Kaneko M, Miyamoto T, Higuchi R (2001) Chem Pham Bull 49:447 168. Yamada K, Sasaki K, Harada Y, Isobe R, Higuchi R (2002) Chem Pham Bull 50:1467 169. Yamada K, Hamada A, Kisa F, Miyamoto T, Higuchi R (2003) Chem Pham Bull 51:46 170. Jeong BY, Ohshima T, Koizumi C (1996) Lipids 31:9 171. Pond DW, Sargent JR (1998) J Plankton Res 20:169 172. Viracaoundin I, Barnathan G, Gaydou EM, Aknin M (2003) Lipids 38:85

Fatty acids from marine I

117

173. Kraffe E, Soudant P, Marty Y, Kervarec N, Jehan P (2002) Lipids 37:507 174. Freites L, Fernandez-Reiriz MJ, Labarta U (2002a) Aquaculture 207: 97 175. Galap C, Netchitaılo P, Leboulenger F, Grillot JP (1999) Comp Biochem Physiol 122A:241 176. Kraffe E, Soudant P, Marty Y, Kervaec N, Jehan P (2002) Lipids 37:507 177. Freites L, Fernandez-Reiriz MJ, Labarta U (2002) Comp Biochem Physiol 132B:453 178. Caers M, Coutteau P, Cure K, Morales V, Gajardo G, Sorgeloos P (1999) Comp Biochem Physiol 123B:97 179. Soudant P, Ryckeghem KV, Marty Y, Moal J, Samain JF, Sorgeloos P (1999) Comp Biochem Physiol 123B:209 180. Garrido JL, Medina I (2002) Analytica Chim Acta 465:409 181. Murphy KJ, Mooney BD, Mann NJ, Nichols PD, Sinclair AJ (2002) Lipids 37:587 182. Abad M, Ruiz, C, Martinez D, Mosquera G, Sanchez JL (1995) Comp Biochem Physiol 110C:109 183. Labarta U, Fernandez-Reiriz MJ, Perez-Camacho A (1999) Comp Biochem Physiol 123A:249 184. Soudant P, Moal J, Marty Y, Samain JF (1997) J Exp Mar Biol Ecol 215:103 185. Brazao S, Morais S, Boaventura D, Re P, Narciso L, Hawkins SJ (2003) Comp Biochem Physiol 136B:425 186. Fernandez-Reiriz MJ, Labarta U, Albentosa M, Perez-Camacho A (1999) Comp Biochem Physiol 124B:309 187. Le Pennec M, Beninger PG, Dorange G, Paulet YM (1991) J Mar Biol Assoc UK 71:451 188. Pazos AJ, Roman G, Acosta CP, Sanchez JL, Abad M (1997) Comp Biochem Physiol 117B:393 189. Le Pennec G, Le Pennec M, Beninger PG (2001) J Mar Biol Assoc UK 81:663 190. Caers M, Coutteau P, Cure K, Morales V, Gajardo G, Sorgeloos P (1999) Comp Biochem Physiol 123B:89 191. Caers M, Coutteau P, Sorgeloos P (1999) Aquaculture 170:307 192. Marty Y, Soudant P, Perrotte S, Moal J, Dussauze J, Samain JF (1999) J Chromatogr A 839:119 193. Jeong BY, Ohshima T, Koizumi C (1999) Comp Biochem Physiol 122B:415 194. de Moreno JEA, Moreno VJ, Ricci L, Roldan M, Gerpe M (1998) Comp Biochem Physiol 119B:631 195. Hayashi K, Kishimura H (2002) J Oleo Sci 51:523 196. Suprayudi MA, Takeuchi T, Hamasaki K (2004) Aquaculture 231:403 197. Takeuchi T, Satoh N, Sekiya S, Shimizu T, Watanabe T (1999) Nippon Suisan Gakkaishi 65:988 198. Kobayashi T, Takeuchi T, Arai D, Sekiya S (2000) Nippon Suisan Gakkaishi 66:1006 199. Hamasaki K, Suprayudi MA, Takeuchi T (2002) Suisan Zoshoku 50:333 200. Suprayudi MA, Takeuchi T, Hamasaki K, Hirokawa J (2002) Suisan Zoshoku 50:205 201. Suprayudi MA, Takeuchi T, Hamasaki, K, Hirokawa, J (2002) Fish Sci 68:1295 202. Lahdes E, Balogh G, Fodor E, Farkas T (2000) Lipids 35:1093 203. Fodor E, Jones RH, Buda C, Kitajka K, Dey I (1995) Lipids 30:1119 204. Cunnane SC (2003) Prog Lipid Res 42:544 205. Wada H, Gombos Z, Murata N (1990) Nature 347:200 206. Miquel M, James D, Dooner H, Browse J (1993) Proc Natl Acad Sci USA 90:6208 207. Meves H (1994) Prog Neurobiol 43:175 208. Xiao YF, Ke Q, Wang SY, Auktor K, Yang Y, Wang GK, Morgan JP, Leaf A (2001) Proc Natl Acad Sci USA 98:3606

118

J.-P. Bergé · G. Barnathan

209. Schmidt A, Wolde M, Thiele C, Fest W, Kratzin H, Podtelejnikov AV, Witke W, Huttner WB, Söling HD (1999) Nature 401:133 210. Wallis JG, Browse J (2002) Prog Lipid Res 41:254 211. Goldberg EM, Zidovetzki R (1997) Biophys J 73:2603 212. Tsutsumi T, Yamauci E, Suzuki E, Watanbe S, Kobayashi T, Okuyama H (1995) Biol Pharm Bull 18:664 213. Wallis JG, Watts JL, Browse J (2002) Trends Biochem Sci 27:467 214. Napier JA (2002) Trends Plant Sci 7:51 215. Connor WE (2000) Am J Clin Nutr 71:171S 216. Albert, CM, Hennekens CH, O’Donnell CJ, Ajani UA, Carey VJ, Willett WC, Ruskin JN, Manson JE (1998) JAMA 279:23 217. Kromhout D (1998) JAMA 279:65 218. Krauss RM, Eckel RH, Howard B, Appel LJ, Daniels SR, Deckelbaum RJ, Erdman JW, Kris-Etherton P, Goldberg IJ, Kotchen TA, Lichtenstein AH, Mitch WE, Mullis R, Robinson K, Wylie-Rosett J, St Jeor S, Suttie J, Tribble DL, Bazzarre TL (2000) Circulation 102:2284 219. O’Keefe Jr, James H, Harris WS (2000) Am J Cardiol 85:1239 220. Oomen CM, Feskens EJ, Räsänen L, Fidanza F, Nissinen AM, Menotti A, Kok FJ, Kromhout D (2000) Am J Epidemiol 151:999 221. Torres IC, Mira L, Ornelas CP, Melim A (2000) Br J Nutr 83:371 222. Iso H, Rexrode KM, Stampfer MJ, Manson JE, Colditz GA, Speizer FE, Hennekens CH, Willett WC (2001) JAMA 285:304 223. Hu FB, Bronner L, Willett WC, Stampfer MJ, Rexrode KM, Albert CM, Hunter D, Manson JE (2002) JAMA 287:1815 224. He K, Rimm EB, Merchant A, Rosner BA, Stampfer MJ, Willett WC, Ascherio A (2002) JAMA 288:3130 225. Calder PC (2003) Ital Heart J 4:427 226. Thies F, Garry JMC, Yaqoob P, Rerkasem K, Williams J, Shearman CP, Gallagher PJ, Calder PC, Grimble RF (2003) The Lancet 361:477 227. Madsen, T, Skou HA, Hansen VE, Fog L, Christensen JH, Toft E, Schmidt EB (2001) Am J Cardiol 88:1139 228. Albert CM, Campos H, Stampfer MJ, Ridker PM, Manson JE, Willett WC, Ma J (2002) N Engl J Med 346:1113 229. Rosenberg IH (2002) New Engl J Med 346:1102 230. Christensen JH, Skou HA, Fog L, Hansen V, Vesterlund T, Dyerberg J, Toft E, Schmidt EB (2001) Circulation 103:651 231. Bigger JT, El-Sherif T (2001) Circulation 103:623 232. De Caterina R, Madonna R, Zucchi R, La Rovere MT (2003) Am Heart J 146:420 233. Dallongeville J, Yarnell J, Ducimetière P, Arveiler D, Ferrières J, Montaye M, Luc G, Evans A, Bingham A, Hass B, Ruidavets JB (2003) Circulation 108:820 234. Marchioli R, Schweiger C, Tavazzi L, Valagussa F (2001) Lipids 36:S119 235. Marchioli R, Barzi F, Bomba E, Chieffo C, Di Gregorio D, Di Mascio R, Franzosi MG, Geraci E, Levantesi G, Maggioni AP, Mantini L, Marfisi RM, Mastrogiuseppe G, Mininni N, Nicolosi GL, Santini M, Schweiger C, Tavazzi L, Tognoni G, Tucci C, Valagussa F, GISSI-Prevenzione Investigators (2002) Circulation 105:1897 236. Leaf A (2002) Circulation 105:1874 237. Lemaitre RN, King IB, (2003) n-3 polyunsaturated FA, fatal ischemic heart disease, and nonfatal myocardial infarction in older adults: the cardiovascular health study. Am J Clin Nutr 77:319 238. Harris WS (2003) Am J Clin Nutr 77:279

Fatty acids from marine I 239. 240. 241. 242. 243. 244. 245. 246. 247. 248. 249. 250. 251. 252. 253. 254. 255. 256. 257. 258. 259. 260. 261. 262.

263. 264. 265. 266. 267. 268. 269. 270. 271. 272. 273. 274.

119

Cobiac L, Nestel PJ, Wing LM, Howe PR (1991) Clin Exp Pharmacol Physiol 18:265 Radack K, Deck C, Huster G (1991) Arch Intern Med 151:1173 Appel LJ, Miller ER, Seidler AJ, Whelton PK (1993) Arch Intern Med 153:1429 Morris MC, Sacks F, Rosner B (1993) Circulation 88:523 Toft I, Bonaa KH, Ingebretsen OC, Nordoy A, Jenssen T (1995) Ann Intern Med 123:911 Connor WE (1995) Ann Intern Med 123:950 Andreassen AK, Hartmann A, Offstad J, Geiran O, Kvernebo K, Simonsen S (1997) J Am Coll Cardiol 29:1324 Bao DQ, Mori TA, Burke V, Puddey IB, Beilin LJ (1998) Hypertension 32:710 Mori TA, Bao DQ, Burke V, Puddey IB, Watts GF, Beilin LJ (1999) Am J Clin Nutr 70:817 Stark KD, Park EJ, Maines VA, Holub BJ (2000) Am J Clin Nutr 72:389 Laidlaw M, Holub BJ (2003) Am J Clin Nutr 77:37 Goodfellow J, Bellamy MF, Ramsey MW, Jones CJ, Lewis MJ (2000) J Am Coll Cardiol 35:265 Flaten H, Hostmark AT, Kierulf P, Lystad E, Trygg K, Bjerkedal T, Osland A (1990) Am J Clin Nutr 52:300 Norrish AE, Skeaff CM, Arribas GL, Sharpe SJ, Jackson RT (1999) Br J Cancer 81:1238 Terry P, Lichtenstein P, Feychting M, Ahlbom A, Wolk A (2001) The Lancet 357:1764 Takezaki T, Hirose K, Inoue M, Hamajima N, Yatabe Y, Mitsudomi T, Sugiura T, Kuroishi T, Tajima K (2001) Br J Cancer 84:1199 Barber MD, Ross JA, Voss AC, Tisdale MJ, Fearon KC (1999) Br J Cancer 81:80 Gogos CA, Ginopoulos P, Salsa B, Apostolidou E, Zoumbos NC, Kalfarentzos F (1998) Cancer 82:395 Kremer JM (2000) Am J Clin Nutr 71:349S Volker D, Fitzgerald P, Major G, Garg M (2000) J Rheumatol 27:2343 Cleland LG, James MJ (2000) J Rheumatol 27:2305 Navarro E, Esteve M, Olivé A, Klaassen J, Cabré E, Tena X, Fernandez-Bañares F, Pastor C, Gassull MA (2000) J Rheumatol 27:298 Darlington LG, Stone TW (2001) Brit J Nutr 85:251 Mayser P, Mrowietz U, Arenberger P, Bartak P, Buchwald J, Christophers E, Jablonska S, Salmhofer W, Schill WB, Krämer HJ, Schlotzer E, Mayer K, Seeger W, Grimminger F (1998) J Am Acad Dermatol 38:539 Danno K, Sugie N (1998) J Dermat 25:703 Hodge L, Salome CM, Peat JK, Haby MM, Xuan W, Woolcock AJ (1996) Med J Aust 164:137 Broughton KS, Johnson CS, Pace BK, Liebman M, Kleppinger KM (1997) Am J Clin Nut 65:1011 Katz DP, Manner T, Furst P, Askanazi J (1996) Nutrition 12:334 Schwartz J (2000) Am J Clin Nutr 71:393S Stevens LJ, Zentall SS, Deck JL, Abate ML, Watkins BA, Lipp SR, Burgess JR (1995) Am J Clin Nutr 62:761 Burgess JR, Stevens L, Zhang W, Peck L (2000) Am J Clin Nutr 71:327S Stordy BJ (2000) Am J Clin Nutr 71:323S Heude, B, Ducimetière P, Berr C, Study E (2003) Am J Clin Nutr 77:803 Edwards R, Peet M, Shay J, Horrobin D (1998) J Affect Disord 48:149 Calabrese, JR, Rapport DJ, Shelton MD (1999) Arch Gen Psychiatry 56:413 Kyle DJ, Schaefer E, Patton G, Beiser A (1999) Lipids 34:S245

120

J.-P. Bergé · G. Barnathan

275. Severus, WE, Ahrens B, Stoll AL (1999) Arch Gen Psychiatry 56:380 276. Stoll AL, Severus WE, Freeman MP, Rueter S, Zboyan HA, Diamond E, Cress KK, Marangell LB (1999) Arch Gen Psychiatry 56:407 277. Bruinsma KA, Taren DL (2000) Nut Rev 58:98 278. Conquer JA, Tierney MC, Zecevic J, Bettger WJ, Fisher RH (2000) Lipids 35:1305 279. Joy CB, Mumby-Croft R, Joy LA (2000) Cochrane Database Syst Rev 4 280. Richardson AJ, Easton T, Puri BK (2000) European Neuropsychopharmacology 10:189 281. Tanskanen A, Hibbeln JR, Hintikka J, Haatainen K, Honkalampi K, Viinamäki H (2001) Arch Gen Psychiatry 58:512 282. Peet M, Horrobin DF (2002) Arch Gen Psychiat 59:913 283. Small MF (2002) New Scientist, August 24:34 284. Levine BS (1997) Nutrition Today 32:248 285. Fidler N, Sauerwald T, Pohl A, Demmelmair H, Koletzko B (2000) J Lipid Res 41:1376–83 286. Olsen S,Secher NJ (2002) Brit Med J 324:1 287. Jensen CL, Maude M, Anderson RE, Heird WC (2000) Am J Clin Nutr 71:292S 288. Makrides M, Gibson RA (2000) Am J Clin Nutr 71:307S 289. Connor WE, Lowensohn R, Hatcher L (1996) Lipids 31:S183 290. Cunnane SC, Francescutti V, Brenna JT, Crawford MA (2000) Lipids 35:105 291. Carlson SE (1999) Acta Paediatr Suppl 88:72 292. Mitchell EA, Aman MG, Turbott SH, Manku M (1987) Clin Pediatr 26:406 293. Uauy-Dagach R, Valenzuela A (1996) Nutr Rev 54:S102 294. Pepping J (1999) Am J Health-Syst Ph 56:719 295. Simopoulos AP (2002) Biomed Pharmacother 56:365 296. de Lorgeril M RS, Mamelle N, Salen P, Martin JL, Mon-jaud I, Guidollet J, Touboul P, Delaye J (1994) Lancet 343:1454 297. Bartram HP, Gostner A, Reddy BS, Rao CV, Scheppach W, Dusal G, Richter A, Richter F, Kasper H (1995) Eur J Cancer Prev 4:231 298. Maillard V, Bougnoux P, Ferrari P, Jourdan M-L, Pinault M, Lavillonnieree FF, Body G, Le Floch O, Chajes V (2002) Int J Cancer 98:78 299. James MJ, Cleland LG (1997) Semin Arthritis Rheum 27:85 300. Heath and Welfare Canada (1990). Table 19. Summary of examples of recommended nutrients based on energy expressed as daily rates. In: Nutrition Recommendations. The reports of the scientific review committee. Health and Welfare Canada, Scientific review committee, Ottawa, 203 (H49-42/1990E) 301. ISSFAL (1999) Adequate intakes. International Society for the Study of FA and Lipids (ISSFAL), Devon (http://www.issfal.org.uk/adequateintakes.htm) 302. BNF (2000) British Nutrition Foundation (BNF), London 303. IOM (2002) Dietary reference intakes for energy, carbohydrates, fiber, fat, protein and amino acids (macronutrients). National Academy of Sciences, food and nutrition board, Institute of Medicine (IOM). National Academy Press (NAP), Washington, DC, 335–432 304. Watkins BA, Lippman HE, Le Bouteiller L, Seifert MF (2001) Prog Lipid Res 40:125 305. Ucciani E (1995) Nouveau Dictionnaire des Huiles Végétales. Compositions en acides gras. Lavoisier TEC&DOC, Paris (France) 306. Abbadi A, Domergue F, Meyer A, Riedel K, Sperling P, Zank TK, Heinz E (2001) Eur J Lipid Sci Technol 103:106 307. Naylor RL, Goldburg RJ, Primavera JH, Kautsky N, Beveridge MC, Clay J, Folke C, Lubchenco J, Mooney H, Troell M (2000) Nature 405:1017

Fatty acids from marine I

121

308. Tocher DR, Leaver MJ, Hodgson PA (1998) Prog Lipid Res 37:73 309. Shanklin J, Cahoon EB (1998) Annu Rev Plant Physiol Plant Mol Biol 49:611 310. Pereira SL, Leonard AE, Mukerji P (2003) Protaglandins, Leukotrienes and essential FA 68:97 311. Napier JA, Sayanova O, Sperling P, Heinz E (1999) Trends Plant Sci 4:2 312. Napier JA, Michaelson LV, Sayanova O (2003) Prostaglandins Leukot Essent Fatty Acids 68:135 313. Satouchi K, Hirano K, Sakaguchi M, Takehara H, Matsuura F (1993) Lipids 28:837 314. Beaudoin F, Michaelson LV, Hey SJ, Lewis, MJ, Shewry PR, Sayanova O, Napier J (2000) Proc Natl Acad Sci USA 97:6421 315. Napier JA, Michaelson LV (2001) Lipids 36:761 316. Watts JL, Browse J (2002) Proc Natl Acad Sci USA 99:5854 317. Parker-Barnes JM, Das T, Bobik E, Leonard AE, Thurmond JM, Chaung LT, Huang YS, Mukerji P (2000) Proc Natl Acad Sci USA 97:8284 318. Gunstone FD (1994) Prog Lipid Res 33:19 319. Khozin I, Adlerstein D, Bigongo C, Heimer YM, Cohen Z (1997) Plant Physiol 114:223 320. Wallis JG, Browse J (1999) Arch Biochem Biophys 365:307 321. Qi B, Beaudoin F, Fraser T, Stobart AK, Napier JA, Lazarus CM (2002) FEBS Lett 510:159 322. Qiu X, Hong H, Mc Kenzie SL (2001) J Biol Chem 276:31 561 323. Voss A, Reinhart M, Sankarappa S, Sprecher H (1991) J Biol Chem 266:19 995 324. Moore SA, Hurt E, Yoder E, Sprecher H, Spector AA (1995) J Lipid Res 36:2433 325. Ferdinandusse S, Denis S, Mooijer PA, Zhang Z, Reddy JK, Spector AA, Wanders RJ (2001) J Lipid Res 42:1987 326. Su HM, Moser AB, Moser HW, Watkins PA (2001) J Biol Chem 276:38 115 327. de Antueno RJ, Knickle, LC, Smith H, Elliot ML, Allen SJ, Nwaka S, Winther MD (2001) FEBS Lett 509:77 328. D’andrea S, Guillou H, Jan S, Catheline D, Thibault JN, Bouriel M, Rioux V, Legrand P (2002) Biochem J 364:49 329. Metz JG, Roessler P, Facciotti D, Levering C, Dittrich F, Lassner M, Valentine R, Lardizabval K, Domergue F, Yamada A, Yazawa K, Knauf V, Browse J (2001) Science 293:290 330. Napier JA, Michaelson LV, Stobart AK (1999) Curr Opin Plant Biol 2:123 331. Cho HP, Nakamura M, Clarke SD (1999) J Biol Chem 274:37 335 332. Leonard AE, Kelder B, Bobik EG, Chuang LT, Parker-Barnes JM, Thurmond JM, Kroeger PE, Kopchick JJ, Huang YS, Mukerji P (2000) Biochem J 347:719 333. Watts JL, Browse J (1999) Arch Biochem Biophys 362:175 334. Michaelson LV, Napier JA, Lewis M, Griffiths G, Lazarus CM, Stobart, AK (1998) FEBS Lett 439:215 335. Michaelson LV, Lazarus CM, Griffiths G, Napier JA, Stobart AK (1998) J Biol Chem 273:19 055 336. Knutzon DS, Thurmond JM, Huang YS, Chaudhary S, Bobik Jr EG, Chan GM, Kirchner SJ, Mukerji P (1998) J Biol Chem 273:29 360 337. Cho HP, Nakamura M, Clarke SD (1999) J Biol Chem 274:471 338. Napier JA, Hey SJ, Lacey DJ, Shewry PR (1998) Biochem J 330:611 339. Sayanova O, Smith MA, Lapinskas P, Stobart AK, Dobson G, Christie WW, Shewry PR, Napier JA (1997) Proc Natl Acad Sci USA 94:4211 340. Sperling P, Lee M, Girke T, Zähringer U, Stymne S, Heinz E (2000) Eur J Biochem 267:3801

122

J.-P. Bergé · G. Barnathan

341. 342. 343. 344. 345. 346.

Girke T, Schmidt H, Zähringer U, Reski R, Heinz E (1998) Plant J 15:39 Yuan L, Kirchner SJ, Mukerji P, Knutzon DS (1999) Lipids 34:649 Sakuradani E, Kobayashi M, Shimizu S (1999) Gene 238:445 Spychalla JP, Kinney AK, Browse J (1997) Proc Natl Acad Sci USA 94:1142 Zank TK, Zähringer U, Lerchl JE, Heinz E (2000) Biochem Soc Trans 28:654 Nichols D, Bowman J, Sanderson K, Mancuso-Nichols C, Lewis T, McMeekin, T, Nichols PD (1999) Curr Opin Biotechnol 10:240 Yazawa K (1996) Lipids 31:S297 Watanabe K, Yazawa K, Kondo K, Kawaguchi A (1997) J Biochem 122:467 Bisang C Long PF, Cortés J, Westcott J, Crosby J, Matharu AL, Cox RJ, Simpson TJ, Staunton J, Leadlay PF (1999) Nature 401:502 Heath RJ, Rock CO (2000) Nature 406:145 Allen EE, Facciotti D, Bartlett DH (1999) Appl Environ Microbiol 65:1710 Tanaka M, Ueno A, Kawasaki K, Yumoto I, Ohgiya S, Hoshino T, Ishizaki K, Okuyama H, Morita N (1999) Biotechnol Lett 21:939 Leonard AE, Bobik EG, Dorado J, Kroeger PE, Chuang LT, Thurmond JM, ParkerBarnes JM, Das T, Huang YS, Mukerji P (2000) Biochem J 350:765 Huang J, Aki T, Yocochi T, Nakahara T, Honda D, Kawamoto S, Shigeta S, Ono K, Suzuki O (2003) Mar Biotechnol 5:451 Müller-Navarra DC, Brett MT, Liston AM, Goldman CR (2000) Nature 403:74 Iida I, Nakahara T, Yocochi T, Kamisaka Y, Yagi H, Yamaoka M, Suzuki O (1996) Journal Ferment Bioeng 81:76 Yokochi T, Honda D, Higashihara T, Nakahara T (1998) Appl Microbiol Biotechnol 49:72 Barclay WR, Meager KM, Abril JR (1994) J Appl Phycol 6:123 Lewis TE, Mooney BD, McMeekin TA, Nichols PD (1998) Chem Aust 65:37 Vazhappilly R, Chen F (1998) Bot Mar 41:553 Singh A, Ward OP (1997) Microbial production of docosahexaenoic acid (DHA, C22:6 ). In: Neidleman SL, Laskin AI (eds.), Advances in Applied Microbiology, Vol 45, pp 271–312 Ratledge C (1998) Opportunities for marine microorganisms for the production of polyunsaturated FA. In: Le Gal Y, Muller-Feuga A (eds.), Marine Microorganisms for Industry. Actes de colloque 21. IFREMER, Plouzane, France, pp 18–25 Lewis TE, Nichols PD, McMeekin TA (1999) Mar Biotechnol 1:580 Ratledge C (1993) Trends Biotechnol 11:278 Circular 886 (Rev 1) 1 Sajbidor J, Dobronova S, Certik M (1990) Biotechnol Lett 12:455 Gandhi SR, Weete JD (1991) J Gen Microbiol 137:1825 Nichols DS, Nichols PD, McMeekin TA (1993) Antarct Sci 5:149 Jøstensen, JP, Landfald B (1997) FEMS Microbiol Lett 151:95 Watanabe K, Sezaki K, Yazawa K, Hino A (1992) Nipp Suis Gakk 58:271 Nichols DS, Hart P, Nichols PD, McMeekin TA (1996) Aquaculture 147:115 Lewis TE, Nichols PD, Hart PR, Nichols DS, McMeekin TA (1998) J World Aquacult Soc 29:313 Gonzalezbaro MD, Pollero RJ (1998) Comp Biochem Physiol 119:747 Erwin J, Bloch K (1964) Science 143:1006 Zhukova NV, Kharlamenko VI, (1999) Aquat Microb Ecol 17:153 DeLong EF, Yayanos AA (1987) Appl Environ Microbiol 51:730 Bowman JP, McCammon SA, Nichols DS, Skerratt JS, Rea SM, Nichols PD, McMeekin TA (1997) Int J Syst Bacteriol 47:1040

347. 348. 349. 350. 351. 352. 353. 354. 355. 356. 357. 358. 359. 360. 361.

362.

363. 364. 365. 366. 367. 368. 369. 370. 371. 372. 373. 374. 375. 376.

Fatty acids from marine I

123

377. Bowman JP, Gosink JJ, McCammon SA, Lewis TE, Nichols DS, Nichols PD, Skerratt JH, Staley JT, McMeekin TA (1998) Int J Syst Bacteriol 48:1171 378. Hugenholtz P, Goebel BM, Pace NR (1998) J Bacteriol 180:4765 379. Yano Y, Nakayama A, Yoshida K (1997) Appl Environ Microbiol 63:2572 380. Kato C (1999) Molecular analysis of the sediment and isolation of extreme barophiles from the deepest Mariana Trench. In: Extremophiles in Deep-Sea Environments, Tsujii K (Ed), Springer-Verlag, Tokyo, pp. 27–38 381. Venkateswaren K, Dollhopf ME, Aller R, Stackebrandt E, Nealson KH (1998) Int J Syst Bacteriol 48:965 382. Ivanova EP, Mikhailov VV (2001) Microbiology 70:10 383. Bozal N, Montes MJ, Tudela E, Jimenez F, Guinea J (2002) Int J Syst Evol Microbiol 52:195 384. Nogi, Y, Kato C, Horikoshi K (1998) J Gen Appl Microbiol 44:289 385. Deming JW, Somers LK, Sraube WL, Swartz DG, MacDonnell MT (1988) Syst Appl Microbiol 10:152 386. Yumoto I, Kawasaki K, Iwata H, Matsuyama H, Okuyama H (1998) Int J Syst Bacteriol 48:1357 387. Urakawa H, Kita-Tsukamoto K, Steven S, Ohwada K, Colwell RR (1998) FEMS Microbiol Lett 165:373 388. Nogi Y, Kato C (1999) Extremophiles 3:71 389. Benediktsdóttir E, Verdonck L, Sproër C, Helgason S, Swings J (2000) Int J Syst Evol Microbiol 50:479 390. Mountfort DO, Rainey FA, Burghardt J, Kaspar HF, Stackebrandt E (1998) Arch Microbiol 169:231 391. Nogi Y, Kato C, Horikoshi K (2002) Int J Syst Evol Microbiol 52:1527 392. Kawasaki K, Nogi Y, Hishinuma M, Nodasaka Y, Matsuyama H, Yumoto I (2002) Int J Syst Evol Microbiol 52:1455 393. Bowman JP, McCammon SA, Lewis TE, Skerratt JH, Brown JL, Nichols DS, McMeekin TA (1998) Microbiology 144:1601 394. Nogi Y, Masui N, Kato C (1998) Extremophiles 2:1 395. Iwanami H, Yamaguchi T, Takeuchi M (1995) Nipp Suis Gakk 61:205 396. Watanabe K, Ishikawa C, Ohtsuka I, Kamata M, Tomita M, Yazawa K, Muramatsu H (1997) Lipids 32:975 397. Barbieri E, Paster BJ, Hughes D, Zurek L, Moser, DP, Teske A, Sogin ML (2001) Environ Microbiol 3:151 398. Ederington MC, Mcmanus GB, Harvey HR (1995) Limnol Oceanogr 40:860 399. Nichols PD, Nichols DS, Lewis T, Bowman JP, Skerrat JH, McMeekin TA (1998) Novel bacteria as alternate sources of polyunsaturated FA for use in aquaculture and other industries. In: Marine microorganisms for industry. Le Gal Y, Muller-Feuga A (eds.), Plouzane: Editions IFREMER, 26–32 400. Russell NJ (1992) Physiology and molecular biology of psychrophilic microorganisms. In: Herbert RA, Sharpe RJ (eds.), Molecular Biology and Biotechnology of Extremeophiles, Blackie, Glasgow, pp 203–224 401. Cronan JE, Rock CO (1996) Biosynthesis of membrane lipids. In: Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology, p 612. Edited by Neidhardt FC, and others. Washington, DC: American Society for Microbiology, 149–160 402. Fujii DK, Fulco AJ (1977) J Biol Chem 252:3660 403. Rawlings BJ (1998) Nat Prod Rep 15:275 404. Siegenthaler PA, Belsky MM, Goldstein S (1967) Science 155:93

124

J.-P. Bergé · G. Barnathan

405. Porter D (1990) Phylum Labyrinthulomycota. In: Handbook of Protoctista. Margulis L, Corliss JO, Melkonian M, Chapman DJ (eds). Boston, Jones and Bartlett, pp 388– 398 406. Bahnweg G, Sparrow FK (1974) Am J Bot 61:754 407. Raghukumar S, Gaertner A (1980) Veroff Inst Meeresforsch Bremerhaven 18:289 408. Raghukumar S (1988) Transcr Br Mycol Soc 90:627 409. Honda D, Yokochi T, Nakahara T, Erata M, Higashihara T (1998) Mycol Res 102:439 410. Naganuma T, Takasugi H, Kimura H (1998) Mar Ecol Prog Ser 162:105 411. Cavalier-Smith T, Allsopp MTEP, Chao EE (1994) Philos Trans R Soc Lond B Biol Sci 346:387 412. Nakahara T, Yokochi T, Higashihara T, Tanaka S, Yaguchi T, Honda D (1996) J Am Oil Chem Soc 73:1421 413. Yaguchi T, Tanaka S, Yokochi T, Nakahara T, Higashihara T (1997) J Am Oil Chem Soc 74:1431 414. Bajpai P, Bajpai BP, Ward OP (1991) Appl Microbiol Biotechnol 35:706 415. Bajpai P, Bajpai BP, Ward OP (1991) J Am Oil Chem Soc 68:509 416. Li ZY, Ward OP (1994) J Indust Microbiol 13:238 417. Singh A, Ward OP (1996) J Indust Microbiol 16:370 418. Singh A, Wilson S, Ward OP (1996) World J Microbiol Biotechnol 12:76 419. Weete JD, Kim H, Ghandhi SR, Wang Y, Dute R (1997) Lipids 32:839 420. Ellenbogen BB, Aaronson S, Goldstein S, Belsky M, (1969) Comp Biochem Physiol 29:805 421. Findlay RH, Fell JW, Coleman NK, Vestal JR, (1986) Biochemical indicators of the role of fungi and thraus-tochytrids in mangrove detrital systems. In: Moss ST (ed.), The Biology of Marine Fungi. Cambridge University Press, Cambridge, pp 91–103 422. Barclay W (1992) US patent 5,130,242 423. Bowles RD, Hunt AE, Bremer GB, Duchars MG, Eaton RA (1999) J Biotechnol 70:193 424. Huang J, Aki T, Hachida K, Yokochi T, Kawamoto S, Shigeta S, Ono K, Suzuki O (2001) J Am Oil Chem Soc 78:605 425. Hammond BG, Mayhew DA, Naylor MW, Ruecker FA, Mast RW, Sander WJ (2001) Regul Toxicol Pharmacol 33:192 426. Barclay WR (1994) US Patent 5,340,742 427. Barclay W, Zeller S (1996) J World Aquacult Soc 27:314 428. Hammond BG, Mayhew DA, Robinson K, Mast RW, Sander WJ (2001c) Regul Toxicol Pharmacol 33:356 429. Hammond BG, Mayhew DA, Naylor MW, Holson JF, Nemec MD, Mast RW, Sander WJ (2001) Regul Toxicol Pharmacol 33:205 430. Fitch Haumann B (1999) INFORM 9:1108 431. Narciso I, Pousao-Ferreira P, Passos A, Luis O (1999) Aquacult Res 30:21 432. Carter CG, Bransden MP, Lewis TE, Nichols PD (2003) Mar Biotechnol 5:480 433. Bull AT, Goodfellow M, Slater JH (1992) Annu Rev Microbiol 46:219 434. Bull AT, Ward AC, Goodfellow M (2000) Microbiol Mol Biol R 46:573 435. Certik M, Shimizu S (1999) J Biosci Bioeng 87:1 436. Becker CC, Kyle DJ (1998) Food Technol 52:68 437. Arts MT, Ackman RG, Holub BJ (2001) Can J Fish Aquat Sci 58:122 438. Sargent JR, Tacon AGJ (1999) P Nutr Soc 58:377 439. Sargent JR (1995) (n-3) polyunsaturated FA and farmed fish. In: Hamilton, RJ, Rice, RD (eds.), Fish Oil: Technology, Nutrition and Marketing. Barnes and Associates, Bucks, pp 67–94

Fatty acids from marine I

125

440. Peng J, Larondelle Y, Ackman RG, Rollin X (2003) Comp Biochem Physiol Pt B 134:335 441. Bell JG, Tocher DR, Farndale BM, Cox DI, McKinney RW, Sargent JR (1997) Lipids 32:515 442. Ackman RG (1980) Fish lipids. In: Connell, JJ (ed.), Advances in Fish Science and Technology. Fishing News, Farnham, p 86 443. Viga A, Grahl-Nielsen O (1990) Comp Biochem Physiol Pt B 96:721 444. Pickova J, Kiessling A, Petterson A, Dutta PC (1999) Fish Physiol Biochem 21:147 445. Ould El Kebir MV, Barnathan G, Siau Y, Miralles J, Gaydou EM (2003) J Agric Food Chem 51:1942 446. Fodor E, Jones RH, Buda C, Kitajka K, Dey I, Farkas T (1995) Lipids 30:1119 447. Lee KH, Jeong ICH, Suh JS, Jung WJ, Kim CG, Lee BH (1986) Han’guk Susan Hakhoechi 19:423 448. Tritar B, Attia-Chaouch S, Hammami M (1997) Ichtyophysiol Acta 20:67 449. Saito H, Ishihara K, Murase T (1997) J Sci Food Agric 73:53 450. Bandarra MN, Batista I, Nunes ML, Empis JM, Christie WW (1997) J Food Sci 62:40 451. Gamez-Meza N, Higuera-Ciapara I, Calderon de la Barca AM, Vazquez-Moreno L, Noriega-Rodriguez J, Angulo-Guerrero O (1999) Lipids 34:639 452. Tanakol R, Yazici Z, Sener E, Sencer, E (1999) Lipids 34:291 453. Marquez-Ruiz G, Velasco J, Dobarganes C (2000) Eur Food Res Technol 211:13 454. Hilbert G, Lillemark L, Balchen S, Højskov CS (1998) Chemosphere 37:1241 455. Njinkoué JM, Barnathan G, Mirallès J, Gaydou EM, Samb A (2002) Comp Biochem Physiol 131B:395 456. Gunstone FD (2002) Market report. Lipid Technology Newsletter 8:5 457. Rees JF, Cure K, Piyatiratitivorakul S, Sorgeloos P, Menasveta P (1994) Aquaculture 122:193 458. Salhi M, Izquierdo MS, Hernandezcruz CM, Gonzalez M, Fernandezpalacios H (1994) Aquaculture 124:275 459. Pike IH, Barlow SM (1999) Fish meal and oil to the year 2010: supplies for aquaculture. Presented at World Aquaculture 1999, Sydney, Australia, April 26–May 2 1999, Abstract, p 603 460. Molyneaux M, Chong ML (1998) Food Technol 52:56 461. Barlow SM (2002) 2nd Seafood By-Products Conference, Alaska, USA 462. Mukherjee KD (1999) INFORM 10:308

Adv Biochem Engin/Biotechnol (2005) 96: 127–163 DOI 10.1007/b135783 © Springer-Verlag Berlin Heidelberg 2005 Published online: 25 August 2005

Fish and Shellfish Upgrading, Traceability Fabienne Guérard1 (u) · Daniel Sellos2 · Yves Le Gal2 1 ANTiOX-UBO,

Pôle universitaire P.J. Helias, Creac’h Gwen, 29000 Quimper, France [email protected] 2 Marine Biology Station, Muséum National d’Histoire Naturelle, BP 225, 29182 Concarneau cedex, France 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Enzymes From Fish and Other Marine Creatures . . . . . . . . . . . . . .

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3 3.1 3.2 3.3 3.4

Fish and Shellfish Protein Hydrolysates . . . . . . . . . . . . . . . From Fish Silage to Fish Protein Hydrolysates . . . . . . . . . . . Advantages of Commercial Exogenous Enzyme Addition . . . . . Quantification of the Proteolysis Extent . . . . . . . . . . . . . . . Mechanism of Hydrolysis and General Properties of Hydrolysates

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131 131 133 137 138

4 4.1 4.1.1 4.1.2 4.1.3 4.1.4 4.1.5 4.2 4.3

Recent Developments in Fish Protein Hydrolysates . . . . . . Biologically Active Substances in By-Product Hydrolysates . . Neuroactive Peptides . . . . . . . . . . . . . . . . . . . . . . . Enzyme Regulators and Inhibitors . . . . . . . . . . . . . . . . Immunoactive Peptides . . . . . . . . . . . . . . . . . . . . . . Hormonal and Hormonal-Regulating Peptides . . . . . . . . . Antioxidant Activities . . . . . . . . . . . . . . . . . . . . . . . Marine Waste as a Nutrient Source in Fermentation Processes Other Applications of FPHs . . . . . . . . . . . . . . . . . . . .

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140 140 140 141 142 142 143 145 146

5 5.1 5.2

Genetic Traceability of Fish and Shellfish Species and By-Products . . . . Choice of Marker Sequences . . . . . . . . . . . . . . . . . . . . . . . . . . Protocols for Fish and Fish Coproduct Identification . . . . . . . . . . . . .

146 147 150

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Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract Recognition of the limited biological resources and the increasing environmental pollution has emphasised the need for better utilisation of by-products from the fisheries. Currently, the seafood industry is dependent on the processing of the few selected fish and shellfish species that are highly popular with consumers but, from economic and nutritional points of view, it is essential to utilise the entire catch. In this review, we will focus on recent developments and innovations in the field of underutilised marine species and marine by-product upgrading and, more precisely, on two aspects of the bioconversion of wastes from marine organisms, i.e. extraction of enzymes and preparation of protein hydrolysates. We will deal with the question of accurate determination of fish species at the various steps of processing. Methods of genetic identification applicable to fresh fish samples and to derived products will be described.

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Keywords Marine by-products · Hydrolysates · Bioactivity · Genetic traceability Abbreviations ACE Angiotensin-converting enzyme CCK Cholecystokinin CGRP Calcitonine gene related peptide CTAB Cetyltrimethylammonium bromide Da Dalton DH Degree of hydrolysis DNA Deoxyribonucleic acid E.C. Enzyme commission nomenclature EDTA Ethyl diamine tetraacetic acid EPB Epoxy-pseudoisoeugenol-(2-methylbutyrate) EtBr Ethydium bromide FP Functional properties FPH Fish protein hydrolysate FPLC Fast protein liquid chromatography GPI Guinea pig ileum test GRAS Generally recognised as safe GRF Growth hormone releasing factor KI Potasium iodide MWDP Molecular weight distribution of peptides N Nitrogen content NaOAc Sodium acetate NB Nitrogen balance NP Nutritional properties NPU Net protein utilisation NR Nitrogen released NSI Nitrogen solubility index OD Optical density OPA o-phtaldehyde Pb Base pair PCR Polymerase chain reaction pH-st pH-stat method RAG-1 Recombination activating gene-1 RFLP Restriction fragment length polymorphism RNA Ribonucleic acid RSM Response surface methodology SC Soluble content SNP Single nucleotide polymorphism STR Short tandemly repeat TAE Tris acetate EDTA TCA Trichloroacetic soluble protein TE Tris-HCl EDTA TFA Trifluoroacetic acid TL Tyrosine level TNBS Trinitrobenzenesulfonic acid TRH Thyrotropin releasing hormone Tris Trishydroxymethylaminomethane

Fish and Shellfish Upgrading, Traceability UF UV

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Ultrafiltration Ultraviolet

1 Introduction In the preparation of sea food products for today’s consumer market, up to 50% of the whole animal is commonly discarded. The remainder is considered as a waste or by-product, even though often high in protein. The wastes from most fishes comprise the skeletal structure, intestinal organs and also a large amount of edible fish muscle that cannot be easily removed from the bone structure of the fish by conventional fish filleting processes. The principle use for the waste has been for fish meal production, with some specific types of waste being utilised for human consumption, e.g. cod livers for cod liver oil [1]. Recognition of the limited biological resources and the increasing environmental pollution has emphasised the need for better utilisation of byproducts from the fisheries. Currently, the seafood industry is dependent on the processing of the few selected fish and shellfish species that are highly popular with consumers, but, from economic and nutritional points of view, it is essential to utilise the entire catch [2]. The increasing demand for protein on a global scale also turns the focus on underutilised protein sources. According to the 2002 FAO annual report [3], the world inland and marine aquatic resources were estimated to be 128.8 million tonnes in 2001. Global production from capture fisheries, aquaculture and the food fish supply is currently the highest on record and remains very significant for global food security, providing more than 15% of total animal protein supplies. Among these resources, marine captures accounted for 86 million tonnes while marine aquaculture accounted for 14.2 million tonnes. This results in about 50 million tonnes of wastes available as a source of raw material including fish guts, which could be further processed for recovering useful enzymes as will be discussed below. Solid wastes generated by seafood plants range from about 30 to 85% of the weight of the landed fish, that is, the portion left after the fillets have been removed, depending upon the type of fishery. Processing of fin fish, crab and shrimp can result in 30–60%, 75–85% and 40–80% waste, respectively. There are many ways in which the fish and shellfish waste could be better utilised, including the following [1]: • Extraction of chitin, enzymes, oils, vitamins, pigments, flavour material • Production of gelatine, chitosan, fish leather Production of novel food ingredients from such underutilised aquatic species or by-products is desirable. According to Shahidi [4], fish proteins possess ex-

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cellent amino acid scores and digestibility characteristics and as such, may be used to enhance the nutritive value of cereal-based foods. In this chapter, we will focus on recent developments and innovations in the field of underutilised marine species and marine by-product upgrading and, more precisely, on two aspects of the bioconversion of wastes from marine organisms, i.e. extraction of enzymes and preparation of protein hydrolysates. We will deal with the question of accurate determination of fish species at the various steps of processing. Methods of genetic identification applicable to fresh fish samples as well as to derived products will be described. The potential uses of marine hydrolysates in the near future may be in the production of bioactive substances as this process is currently under control, in the case of casein and soya hydrolysates. We will not deliberately discuss fish meal for animal nutrition, fish protein concentrates as a cheap nutritious protein source for developing countries, and fish silage product development using mince from low-cost fishery resources (e.g. sausages, etc.) (see the reviews of Sikorski & Naczk [5], Raa & Gildberg [6], Venugopal and Shahidi [7], respectively).

2 Enzymes From Fish and Other Marine Creatures In recent years, a number of enzymes from fish processing wastes have become commercially available for food and other applications, as reviewed by Haard [8]. Aquatic organisms include a wide and extensive taxonomic diversity and many organisms occupy unusual environmental habitats, thus conferring to enzymes some unique characteristics such as psychrophilic properties. The most extensively studied enzymes from the marine environment include pepsin, trypsin, chymotrypsin, elastase, collagenase and alkaline phosphatase isolated from Atlantic cod (Gadus morhua) viscera [9– 11], polar cod (Boreogadus saida) [12], dogfish [13–15], salmon [16, 17] and tropical tuna [18]. Several of these enzymes from poikilothermal organisms are cold-active and have a catalytic activity equal or higher than mammalian enzymes. For example, the temperature optimums of trypsin and alkaline phophatase from cold-water fish are about 30 ◦ C lower than the homologues from warmwater fish or mammals [19, 20]. This property is advantageous in applications where it is desirable to inactivate the enzyme with a mild heat treatment [8]. Nevertheless, a trypsin purified from the pyloric caeca of white craker (Micropogonias furnieri) exhibited a temperature optimum of 60 ◦ C. This could be related to the warm water in which the fish lived [21]. Enzymes like LDH from deep-water fish that live at high pressure have a tighter polypeptide structure than homologues at normal atmospheric temperature, thus making them

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more resistant to proteolytic degradation and more suitable for applications where proteases might interfere with their activity [22]. The gastric proteases of fish that take in saltwater during feeding are salt-activated, in contrast to homologues from mammals that are inhibited by NaCl. This property may be advantageous in applications, such as fermentations, silage and fish sauce, where significant amounts of salt are present [23]. Lysozyme from Artic scallop [24] and other fish have a unique ability to attack both Gram-positive and negative bacteria. According to Vilhelmsson [25], research in this area dates back at least to the early 1970s and the literature was sparse on the subject. However, there has recently been a surge of interest in enzymes from the marine environment and their potential use in food processing, mainly in North European countries. These include a commercially available collagenase preparation from crab hepatopancreas that may have several applications, such as the deskinning of squid (Loligo sp.,), production of caviar, and ripening of salt fish. Cold-active fish pepsins from species such as Atlantic cod (G. morhua) and orange roughly (Hoplostetus atlanticus) have been used for caviar production from the roe of various species. These proteolytic enzymes are used to ease the riddling process, increasing the yield from 70% to 90% for salmon (S. salar) for example. A protease preparation from Novo Nordisk was found to double the yield of roe from rainbow trout O. mykiss [26]. In conclusion, these examples illustrate the fact that enzymes from aquatic organisms will never fit more than a niche in food processing because of the cut-throat competition with enzymes of microbial origin. However, with the advent of recombinant DNA technology, there is also growing interest in cloning genes for unique biochemicals from exotic aquatic organisms for mass production by microbial or other expression systems [8].

3 Fish and Shellfish Protein Hydrolysates 3.1 From Fish Silage to Fish Protein Hydrolysates Fish silage is a liquid product, made from whole fish or parts of fish, to which no material has been added other than a mineral acid to lower the pH to values below 4.5. Liquefaction is carried out by endogenous enzymes naturally present in the fish. Acid aids in accelerating the process by creating the right conditions for the enzymes to work and by helping to break down bone. This procedure efficiently prevents growth of spoilage bacteria [6]. After liquefaction, it is convenient to remove the oil coming from the raw material. The protein in the aqueous layer may thereafter be dried or semi-dried. The main advantages of fish silage are the recovery of fish offal and waste fish, low cost, good nutritional

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value of the resulting product and long storage life. The main inconvenience is the impossibility of regulating the degree of hydrolysis achieved. In recent years the enzyme-catalysed process of hydrolysis, as applied to protein-containing raw materials used by the food industry, has been the object of numerous studies. It has been found that hydrolysis of the proteins themselves may increase yields in recovery processes, improve functional properties or improve process methodology, e.g. with regards to possible means of control. In more general terms, in the mild conditions that characterise enzymatic processes, a protein tends to retain its nutritive value better than in traditional acidic or alkaline hydrolysis [27]. Fish protein hydrolysates (FPHs) are products with high protein content and a wide variety of uses, as reviewed by Mackie [28] and Kristinsson and Rasco [29]. There are several methods of FPHs production, which include utilisation of acids, bases [30], endogenous enzymes or exogenous proteases. Research during the past 20 years has greatly increased the understanding of how to process fish or shellfish by-product hydrolysates [25, 31–34]. Generally, underutilised fish, fish frames or crustacean wastes are suspended in water and enzyme is added to the slurry. In some cases, the meat is first heated in order to denature the endogenous proteases [35]. The reaction is allowed to proceed from under 1 h to 1 week, depending on the activity of the enzyme employed, process temperature and other factors. After separation of solids, pH is adjusted and the aqueous layer is clarified, and then dehydrated. Figure 1 outlines the main steps of the production of protein hydrolysates from the raw material. Cassia et al. [36] described the obtention of FPHs using an autolytic process. The authors concluded that enzymatic autolysis might be a simple and efficient process for upgrading fish filleting wastes. However, although the FPHs had high protein and low lipid content, together with an amino acid composition similar to FAO/WHO standards, the process yield was rather low (< 7%). Shahidi et al. [4] showed that endogenous enzyme alone produced hydrolysates with a protein recovery of approximately 23%, whereas a yield of 51.6–70.6% was obtained for commercial enzymes. Shahidi & Synowieck [30] described an alkali-assisted extraction of proteins from meat and bone residues of harp seal with a high recovery of proteins, ranging from 57% to 64%, together with a high level of taurine, excellent emulsifying capacity and emulsion stability. In addition, processing by-products from shellfish is made up of protein residues from body sections such as heads, carapace and exoskeleton. Enzyme-assisted proteolysis of shellfish processing discards may be used to recover the chitin and nutritionally valuable protein hydrolysate containing up to 64% of protein and 81% of total nitrogen in the product [37], or to extract proteins with flavour-enhancing effects, including carotenoids and/or carotenoproteins [38]. A new process for advanced utilisation of shrimp wastes that includes enzymatic hydrolysis was recently described. The authors

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Fig. 1 Flowsheet for the enzymatic hydrolysis of fish or shellfish proteins to make fish or shellfish protein hydrolysates

demonstrated the recovery of amino acids, nitrogen and astaxanthin by Alcalase pre-treatment of shrimp waste before further processing in chitosan. The nitrogen recovery was about 70% as compared to only 15% by conventional methods. The yield and quality of chitosan was not affected by the enzymatic treatment. In addition, a concentrate of astaxanthin was recovered and could constitute a valuable supplement in salmon feed, improving both the growth and the disease resistance of the fish [39]. 3.2 Advantages of Commercial Exogenous Enzyme Addition The solubilisation of fish tissue in traditional silage production is a timeconsuming process. After 3–10 days, depending on the storage temperature, the degree of hydrolysis (DH) is around 20–70%. Addition of commercial exogenous enzymes to the fish tissue reduces the time needed to obtain a similar DH and allows a control of the DH, and subsequently of the peptide size obtained. The choice of hydrolysis process will depend on the targeted applications. For dietary use, or in order to obtain a hydrolysate with a high nutritional and therapeutic value, it has been shown that protein hydrolysates should be rich in low molecular weight peptides, with as few free amino acids as possible [40]. On the other hand, large molecular weight peptides (more than 20 amino acid residues) are presumed to be associated with the improvement functionality of hydrolysates.

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Table 1 Comparison between chemical and enzymatic hydrolysis [41, 42, 48]

Acid/ alkaline hydrolysis

Enzymatic hydrolysis

Specificity

Advantages

Disavantages

Random process

Fast reaction Complete hydrolysis

High temperatures Molecular weight out of control

Low cost

Large amount of salt

High solubility

Undesirable side reactions (destruction of tryptophan, racemisation, etc.)

Control of the molecular weight

Higher cost

Digestion under mild conditions

Subsequent deactivation of the enzyme

Attractive functional product characteristics

Time consuming

Unique specificity

Few side reactions No destruction of amino acids Higher nutritional value

Enzymatic hydrolysis using commercial exogenous proteases presents a lot of advantages compared to chemical hydrolysis (Table 1). Most commercial proteases can be used to solubilise marine wastes. They are obtained from animal viscera, plants and GRAS microorganisms. Selected examples of proteolytic enzymes used to hydrolyse marine by-products, are presented in Table 2. With regards to the effect of the concentration of enzyme in the reaction solution, it has been found that the percent hydrolysis increases with higher concentrations of enzyme, but only up to a certain point. When using an endopeptidase, such as Alcalase 2,4 L for hydrolysis, so as to moderate DH values (i.e. below DH 20%), the relative content of free amino acids and dipeptides is likely to be low [43]. By selecting both the enzyme and the conditions of digestion, various degrees of hydrolysis or breakdown of the proteins can be achieved in order to obtain products with a range of functional properties. Much of the work is still at the laboratory or pilot-scale level, but there is good reason to be confident that these biological processes will make some of this “waste” protein available as hydrolysates containing bioactive substances (see Sect. 4). In some cases, experimental designs were employed to optimise hydrolysis conditions. Simpson et al. [56] used a 3 × 3 factorial central composite

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Table 2 Selected examples of proteolytic enzymes used to hydrolyse marine wastes Enzymes Suppliers Substrates

Applications

Evaluation of hydrolysis

References

Papain

Solvay Herring Enzymes (Clupea harengus)

1

TCA soluble N, total N, color, sensory, MWDP

[44]

Sigma

1

TL, NP, NSI

[31, 45]

1, 2

pH-st, FP, NP

[4, 46]

1

pH-st, NR

[33]

1

TL, NP, NSI

[31, 45]

1

pH-stat, MWDP

[46]

1

TL, NP, NSI

[31, 45]

1, 2

pH-st, FP, NP

[4, 32, 46]

1

pH-stat, NP

[37]

1

pH-stat, NSI, FP

[48]

1, 2

pH-stat, MWDP

[35, 49]

1

TCA soluble N, [44] total N, colour, sensory, MWDP DH-α-amino acid, [50] NR, colour pH-stat, RSM,NR [34, 51]

Sigma Merck Pepsin

Sigma Sigma

Fungal Sigma protease type II from A. oryzae Alcalase Novo 2, 4 L Nordisk

Lobster cephalothorax (Palinurus sp.) Capelin (Mallotus villosus) Sardine (Sardina pilchardus) Lobster cephalothorax (Palinurus sp.) Atlantic cod (Gadus morhua) Lobster cephalothorax (Palinurus sp.)

Capelin (Mallotus villosus) Shrimp wastes (Crangon crangon) Salmon muscle (Samon salar) Tuna stomach (Tunus albacora) Herring (Clupea harengus)

Pacific whiting 1 (Merluccius productus) Dogfish fillet np (Squalus acanthias) Atlantic cod 1 (Gadus morhua) and salmon (Salmo salar) Shrimp waste 1 (Pandalus borealis) Sardine 1, 2 (Sardina pilchardus) Harp seal 1,2 (Phoca groenlandica)

pH-stat, MWDP

[47]

NR

[39]

pH-stat, MWDP

[52]

pH-stat, FP

[4, 53]

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Table 2 (continued) Enzymes

Suppliers Substrates

Alcalase 0, 6 L Neutrase 0.5 L

Novo Nordisk Novo Nordisk

PTN 3.0 type special Corolase 7089, Corolase PN-L, Flavourzyme 1000 L Umamizyme Protamex

Novo Nordisk Novo Nordisk

Novo Nordisk

Applications

Sardine 1 (Sardina pilchardus) Capelin 1, 2 (Mallotus villosus) Sardine 1 (Sardina pilchardus) Pacific whiting 1 (Merluccius productus) Harp seal 1 (Phoca groenlandica) Capelin 1 (Mallotus villosus) Salmon muscle 1 (Salmo salar)

Tuna stomach (Tunus albacora) Frames of Atlantic Salmon (Salmo salar L)

1 np

Evaluation of hydrolysis

References

pH-st, NR

[33]

pH-st, FP, N,

[4, 32, 46]

pH-st, NR

[33]

DH-α-amino acid, NR, colour pH-stat, FP

[50]

[4, 53]

pH-st

[32]

pH-stat, NSI, FP

[48]

pH-stat, MWDP NR

[54] [55]

1 dietary protein source, 2 biological activities, np not precised, TL tyrosine level, pHst pH-stat method, SC soluble content, N nitrogen content, TCA TCA soluble protein, FP functional properties, NP nutritional properties, NSI nitrogen solubility index, NR nitrogen released, RSM response surface methodology, MWDP molecular weight distribution of peptides

design to optimise hydrolysis of shrimp for recovery of amino acids. The second order polynomial models they proposed could be used to predict the content of specific amino acids to a reasonable degree of accuracy. The response surface regression procedure of the statistical analysis system was used by Shahidi et al. [46, 53] in order to fit a quadratic polynomial equation to the experimental data. The three-dimensional response surface indicated that both the Alcalase concentration and the treatment temperature affected the DH and thus the protein recovery. Response surface methodology was used by several authors in order to study the effects of pH, temperature, enzyme/substrate ratio and substrate concentration on the degree of hydrolysis of crayfish by-products [57] and dogfish muscle [51, 58]. The resulting equations were adequate for predicting the DH under any combination of values of the variables.

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3.3 Quantification of the Proteolysis Extent When using exogenous proteases, the hydrolysis reaction must be carefully controlled in order to maintain uniform quality of the end products. The hydrolysis degree (DH), which is defined as the percentage of cleaved peptide bonds, serves as the controlling parameter for the hydrolysis reaction. The pH-stat technique consists in adding acid or base in order to titrate the released α-amino and α-carboxyl groups, thus maintaining the pH constant. Equation 1 relates the DH to alkali consumption [59]: DH = B × Nb

1 1 1 100% α MP htot

(1)

where α–1 is the calibration factor for the pH-stat, and is the reciprocal of the degree of dissociation: 10pH-pK (2) 1 + 10pH-pK The principle of the pH-stat method has been used by many workers for kinetic studies and in order to monitor the degree of hydrolysis attained by enzymatic reactions on food proteins. In addition to the pH-stat method, the extent of proteolysis may also be quantified by the depression of the freezing point, which is indicative of the increasing osmolarity (osmometry), or by the increase in solubility in trichloracetic acid. DH values determined by different methods are often not directly comparable. Base consumption and osmometry methods are easy to perform, allowing continuous monitoring of the hydrolysis process, whereas the estimation of soluble-nitrogen content using the Kjeldahl method is timeconsuming and cannot be used as an on-line process control tool [60]. The trinitrobenzenesulfonic acid (TNBS) method developed in 1979 by AdlerNissen [61] is always used to determine the DH of food protein hydrolysates. However, the o-phtaldehyde (OPA) method used for analysing the protein hydrolysate DH, has been found to be more accurate, easier and faster to carry out than the TNBS method [62]. This method has a broader application range and is environmentally safer. In addition, Silvestre [63] proposed a review of various methods used for the analysis of protein hydrolysates and discussed the potential and limitations of the different techniques. Finally, hydrolysates can be characterised according to the peptide size in order to check that hydrolysates can be produced in a repeatable manner. In this case, size-exclusion chromatography is a simple and quick method for the evaluation of peptide molecular weight. Gel chromatography of protein hydrolysates is generally performed on a filtered 5 or 10% solution of the hydrolysate in eluent. Separations are carried out on Sephadex G-50 gel or Biogel P-10, which have the nominal working range for protein and peptides α=

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of 1500–30 000 Da and 1500–20 000 Da, respectively [36, 44]. New size exclusion chromatography supports in FPLC mode are now available such as the Superdex Peptide HR 10–30 and Superdex (Pharmacia Biotech, Sweden), with fractionation range from 100 to 7000 Da and 300 to 30 000 Da respectively. Acetonitrile is used as the mobile phase (acetonitrile/distilled water by volume ranging from 2 : 8 to 3 : 7, with 0.1% trifluoroacetic (TFA) [35, 55]. The chemiluminescent nitrogen detection coupled with size exclusion chromatography is also a useful technique for estimating the average molecular weight of peptides and for characterising protein hydrolysates [64]. In practice, the fractionation range values can only serve as guidelines, especially because the elution behaviour of peptides in non-dissociating media is influenced by adsorption and aggregation [43] and because of the underestimation of small peptides and free amino acids [49]. 3.4 Mechanism of Hydrolysis and General Properties of Hydrolysates Under optimal conditions of digestion by the enzyme, fish tissues are converted rapidly from a viscous mince to a free-flowing liquid. Although the detailed mechanism is not understood, it is believed that, on adding fish mince to the suspension, the enzyme is absorbed onto the suspended particles. Simultaneously, hydrolysis of the enzyme-sensitive peptide linkages takes place. The hydrolysis is characterised by an initial rapid phase, during which a large number of peptide bonds are hydrolysed. Subsequently, the rate of hydrolysis decreases and enters a stationary phase with no apparent hydrolysis taking place (Fig. 2). Such kinetics are typical of fish protein hydrolysis, regardless of the enzyme or fish being used. It is also a feature of such processes that approximately 20% of the total nitrogen remains insoluble even when further amounts of enzyme are added during the stationary phase of hydrolysis. To some extent this is believed to be product inhibition, as higher yields of soluble protein can be obtained by reducing the concentration of solids [28, 46]. The nutritional value of FPH is determined both by utilisation of (i) several data such as the proximate composition of the FPH (crude protein, dry matter, ash, amino acid composition) and (ii) by experiments with living animals such as rats, with the calculation of various indexes such as net protein utilisation (NPU), nitrogen balance (NB) and digestibility. In addition, for the FPH to be of a high nutritional value, it has been shown that the dietary proteins should be rich in low-molecular-weight peptides, especially di-and tripeptides, with amounts of free amino acids as low as possible [40]. The nutritional value of a FPH made from cod frames hydrolysed by Alcalase (150 min) and followed by treatment with Kojizyme (510 min), was established in an experiment with rats. The apparent digestibility was significantly (p < 0.05) higher for the group receiving diets in which all the dietary

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Fig. 2 Effect of enzyme/substrate ratio (w/w protein) ranging from 0.1% to 1.5% on the degree of hydrolysis. Substrate is tuna stomach and enzyme is Umamizyme [54]

proteins come from the FPH as compared to rats fed with either none or the lowest inclusion levels of FPH. However, the nitrogen balance (that measures the proportion of nitrogen retained within the body) was significantly higher for the group receiving 10% FPH as compared to rats fed higher inclusion levels [47]. The hydrolysates prepared by Shahidi [4] from capelin (Biocapelin) and seal meat (Bioseal-L) have excellent solubility characteristics at pH values ranging from 2.0 to 10.4. While 90.36–98.57% of nitrogenous compounds of Biocapelin were soluble, corresponding values for Bioseal-L varied between 93.45% and 98.05%. The fat adsorption, moisture retention, emulsification properties and whippability of the hydrolysates were also excellent. The amino acid composition of hydrolysates obtained from the cod filleting waste is closely similar to that of fillets, except for glycine and proline amounts, whose highest concentrations are derived from the relatively higher amount of connective tissue proteins [28, 65]. Most of hydrolysates, however, have varying degrees of a bitter flavour which, although they are mild compared with those of casein hydrolysates, does restrict their application. Bitter tastes are a common feature of enzymically produced protein hydrolysates and are believed to be due to low molecular weight peptides (up to 6000 Da), with hydrophobic side chains normally located in the interior of an intact protein. Until now, the bitter taste was reduced, but not eliminated, by controlling the degree of hydrolysis and the choice of enzyme preparation, so that predominantly tasteless larger molecular weight peptides are produced [47].

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4 Recent Developments in Fish Protein Hydrolysates 4.1 Biologically Active Substances in By-Product Hydrolysates A large number of biologically active peptides have been isolated from bacterial, fungal, plant and animal sources or generated from proteins by enzymatic hydrolysis [66–70]. These peptides, which are inactive within the sequence of the parent protein, can be released by enzymatic proteolysis, for example during gastrointestinal digestion or during food processing. Peptides derived from milk proteins deserve special attention. A wide range of physiological activities including opioid, hypotensive (anti-ACE), immunomodulating, antithrombic, antimicrobial, antiviral and mineral absorption regulatory functions, have been associated with the specific sequences derived from milk (for reviews, see Meisel [71, 72], Clare and Swaisgood [73]; Clare et al. [74]; Floris et al. [75]). In addition, many milk-derived peptides revealed multifunctional properties, i.e. specific peptide sequences having two or more different biological activities such as opioid, ACE-inhibitory and immunomodulatory effects [71]. These bioactive peptides have been characterised in detail. Generally, these structures usually contain 3–20amino acid residues per molecule. Some of them are often further modified through glycosilation, phosphorylation, and/or acylation of multifunctional amino acid residues [66]. The isolation and characterisation of new biologically active peptides is ongoing and will undoubtedly continue in the future. This is partially due to increased knowledge of enzymes and food proteins (inherent amino acid composition and sequence). Consequently, some interesting and very promising new applications for the FPHs have emerged and, at present, a few examples of peptides exhibiting various activities (e.g. opiate, antithrombic or antihypertension activity, immunomodulation, antioxidant) have been reported. The objective of this section is to provide the reader with an up-to-date summary and analyse the new trends and ideas that have emerged in the last few years in this rapidly developing area. 4.1.1 Neuroactive Peptides The endogenous opioids, enkephalins, endorphins and other neuroactives such a somatostrain, bradykinin and thyrotropin-releasing hormone (TRH) are a few examples of neuroactive peptides. These peptides play fundamental roles in the functioning of the nervous system such as in the regulation of pain perception [66].

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Opioid receptors are located in the nervous, endocrine and immune systems, as well as in the tract of the mammalian organism and can interact with their endogenous ligands, as well as with exogenous opioids and opioid antagonists. It has been shown that a variety of neuroactive peptides are formed on the hydrolysis of milk, soy, cereal and fish proteins (for review, see Schlimme & Meisel [76]). For example, opioid peptides such as enkephalins have an affinity for opiate receptors as well as opiate-like effects, inhibited by naloxone. Most of the typical opioid peptides have the same N-terminal sequence, Tyr-Gly-Gly-Phe. Opioid peptides exert their activity by binding to specific receptors of the target cell. Recently, some “pseudo” opioid activities were found in shrimp and cod head hydrolysates, since the inhibition of the contractions measured in the GPI test was naturally reversed without the help of naloxone [77]. In addition, fish protein hydrolysates commonly used as nutritional supplements (commercial names PC60 and Stabilium 200) were reported to reduce anxiety in humans and to improve memory and learning performances in rats and patients [78–80]. PC60 was compared to a potent anxiolytic drug – diazepam (Valium), which acts on benzodiazepine receptors, confirming the anxiolytic properties of the nutritional supplement previously reported in both rats and humans [81]. 4.1.2 Enzyme Regulators and Inhibitors A very important group of peptides is the angiotensin-converting enzyme (ACE) inhibitors, which are currently in use as antihypertensive agents. Hypertension is a major risk factor in cardiovascular disease such as heart disease and heart stroke. Angiotensin I-converting enzyme (ACE, peptidyl dipeptide hydrolase, EC 3.4.15.1.) has been classically associated with the renin-angiotensin system, which regulates peripheral blood pressure. ACE may modulate blood pressure by catalysing the conversion of pro-peptide angiotensin I to a potent vasoconstrictor peptide angiotensin II. Consequently, ACE-inhibitors may exert an inhibitory effect. Moreover, inhibition of ACE may influence different regulatory systems of the organisms involved in modulating blood pressure, immune defence and nervous system activity [71]. These peptides often contain proline, lysine or arginine as the Cterminal residue and have been found in the enzymatic hydrolysate of bovine and human caseins [82–84], zein protein hydrolysates [85] and other food proteins. Peptides obtained from enzymatic hydrolysis of tuna muscle [86, 87], sardine muscle [88], bonito bowels [89] and squid liver and mantle muscle [90], were found to have potent inhibitory effects on ACE.

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In addition, some purified fractions of an Alcalase sardine by-product hydrolysate gave inhibition indexes reaching 90% [77] and may provide a potential good source of antihypertensive molecules. A cod frame hydrolysate was separated, based on the molecular weight of the peptides, through a series of UF-membranes with molecular cut-offs ranging from 30 to 3 kDa. The 3-K hydrolysate had excellent ACE inhibitory activity, while the 10-K showed high antioxidative activity [91]. Until now, antihypertensive effects in humans have not been proven for most of the peptides obtained by processing food proteins. However, recent studies provided proof of antihypertensive effect of the peptides Val-Pro-Pro, Ile-Prp-Pro and Val-Tyr ingested with milk twice a day for several weeks by hypertensive patients in Japan and Finland [92–94]. 4.1.3 Immunoactive Peptides The bioactivity of immunopeptides is characterised by different in vitro and in vivo tests. Many immunomodulating peptides that stimulate proliferation of human lymphocytes and phagocytic activities of macrophages were isolated from milk caseins. For example, Kayser & Meisel [95] reported that the immunoreactivity of human peripheral blood lymphocytes was either stimulated or suppressed by various bioactive peptides derived from milk. A pepsin-chymosin digest of bovine casein induced a significant proliferative response in rat lymphocytes [96]. The presence of immunomodulating effects was demonstrated in various fish hydrolysates. For example, hydrolysates from Atlantic cod, Gadus morhua L., were used both in vitro and in vivo in stimulatory experiments with head kidney leucocytes from Atlantic salmon (Salmo salar). The authors concluded that acid peptide fractions from fish protein hydrolysate may be useful as adjuvants in fish vaccine and as an immune stimulant in fish feed [97, 98]. 4.1.4 Hormonal and Hormonal-Regulating Peptides Selected examples of activities identified in marine waste hydrolysates are the peptides gastrin, growth hormone releasing factors (GRFs), calcitonin and CGRP. These short peptides often exert complex and multiple physiological effects by serving as hormones themselves and/or regulating hormonal responses associated with the control of important metabolic, growth and development processes [65]:

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• Calcitonin gene related peptide (CGRP). This 37 amino acid neuropeptide is derived from the same gene as calcitonin by a mechanism of alternative splicing. It is predominantly synthesised in neural tissue and is mainly involved in the control of vasodilatation, with inotropic and chronotropic effects on the heart [99], but is also involved in the regulation of gastric acid secretion [100]. This peptide can also inhibit the proliferative response of T lymphocytes to mitogens [101] and macrophage activation [102]. In addition, at high doses, CGRP induces the same effect as calcitonin, that is hypocalcemia and hypophosphatemia [103]. The presence of CGRP immunorelated molecules was demonstrated in various fish hydrolysates (cod, sardine, hake and tuna Alcalase 2,4 L-assisted hydrolysates) [104]. CGRP-related molecules were purified from sardine hydrolysates prepared using 0.1% Alcalase and 2 h hydrolysis. Twentytwo mg of crude extract yielded 14 µg of CGRP-related molecules with a purification factor of 12 500. The molecular weight determined by mass spectrometry was 6000 Da. The purified molecules induced an inhibition of the CGRP-stimulated adenylate cyclase activity. This effect was specific as no such effect was observed on the glucagon- stimulated adenylase cyclase activity measured in the same rat liver membrane preparation, suggesting that the purified molecules may act as antagonists for peptides that bind to CGRP receptors in rat liver membranes [105]. • Cholecystokinins (CCK) and gastrins. These belong to a family of short peptides exhibiting a large spectrum of activities including mediation and stimulation of protein synthesis, control of intestinal mobility and secretion of digestive enzymes. Hydrolysates derived from fish heads and viscera (sardine and cod) and shrimp wastes presented a positive response to gastrin radioimmunoassays, thus suggesting the presence of gastrin-like and cholecystokinin-like peptides [106, 107]. In addition, Alcalase 2,4 Lassisted hydrolysis of tuna, shrimp, cod heads and cod muscle significantly stimulated the growth of 3T3 fibroblastic cells suggesting the presence of growth factor-like molecules in the hydrolysates [49, 52]. 4.1.5 Antioxidant Activities In addition to the biological activities described above, another promising research area for the coming years is the presence of antioxidant compounds in marine hydrolysates. The term antioxidant is defined as “any substance that, when present at low concentrations compared to that of an oxidisable substrate, significantly delays or inhibits oxidation of that substrate” [108]. Antioxidants can act at different levels in an oxidative sequence. This may be illustrated by considering one of the many mechanisms by which oxidative stress can cause damage by stimulating the free radical chain reaction of lipid peroxidation. Free radical chain reactions within a material may be inhibited

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either by adding chemicals that retard the formation of free radicals (preventive antioxidants), or by introducing substances that compete for the existing radicals and remove them from the reaction medium (chain breaking antioxidants). Synthetic antioxidants such as 3-ter-butyl-4-hydroxyanisole (BHA), 3,5-di-tert-butyl-4-hydroxytoluene (BHT), tertiary-butylhydroxyquinone and propyl galate are used as food additives to retard lipid oxidation. However, use of synthetic antioxidants in food products is under strict regulation due to the potential health hazards caused by such compounds. Therefore, search for natural and safer antioxidants as alternatives to synthetic ones is of great interest among researchers. Several studies have described the antioxidative activity of proteins and protein hydrolysates such as milk casein, soybean protein, broad beans, bovine serum albumin, wheat gliadin and sunflower meals [109–112]. For example, a hexapeptide with strong free radical scavenging activity was separated from casein hydrolysate. Six antioxidative peptides were isolated from the hydrolysate of a soybean protein, β-conglycinin. These peptides were composed of 5–16 amino acid residues and included hydrophobic amino acids, Val and Leu, at the N-Terminus and Pro, His, or Tyr in their sequences [113]. Two peptides composed of 10 and 15 amino acid residues, and both containing a leucine residue at their N-terminal position, were purified from an Alcalase digest of a by-product of lecithin extraction from egg yolk [108]. A few antioxidant compounds were also isolated from marine hydrolysates such as mackerel (Scomber australasicus) [114], capelin (Mallotus villosus) and harp seal (Phoca groenlandica) [115, 116], and cod frames [91]. Kim et al. [117] isolated two peptides from Alaska pollack skin, composed of 13 and 16 amino acid residues. Both peptides contained a Gly residue at the C-terminus and the repeating motif Gly-Pro-Hyp. In addition, antioxidant compounds were purified from shrimp shell wastes and rockfish [118]. Some of these compounds were identified using mass spectrometry. One antioxidant was proposed to be 1,2-diamino-1-(ohydroxyphenyl)propene [119]. Three antioxidant peptides were isolated and identified from a pepsin digest of prawn (Penaeus japonicus) muscle [120]. Guérard et al. [121] hydrolysed shrimp wastes using Alcalase 2,4 L. By combining membrane filtration separation and chromatography techniques, the antioxidant fraction was partly purified and its molecular weight was estimated to range from 300 to 400 Da. An oyster (Crassostera gigas) extract was prepared from fresh raw oysters and demonstrated a high free radical scavenging activity towards superoxide and hydroxyradical. However, the authors did not establish which compounds in this extract were responsible for the scavenging effect on the radicals [122] Maillard reaction products may also show antioxidant properties derived from their ability to bind heavy metals (particularly iron), thereby giving rise to oxidatively inactive complexes [123]. For example, when reacting a tuna

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stomach hydrolysate with glucose, the antioxidant effect evaluated using the β-carotene-linoleate system model, was increased by 20 to 30% [124]. To summarise this section, all the bioactive substances have been studied by means of the following investigation techniques: • Establishment of an in vitro assay system to determine the biological activity • Hydrolysis of proteins by proteases • Partial isolation of peptides and, sometimes, purification and determination of the structure • In a few cases, synthesis of peptides for the verification of activity In the final report of the European research program FAIR CT 97-3097 (acronym HYDROFISH), numerous examples of biological activities identified in fish and shrimp waste hydrolysates are presented. In conclusion to this section, marine hydrolysates or extracts, particularly from fish viscera or marine invertebrates do contain various biological activities that should be further investigated. Remarkable observations have already been made during feeding with various extracts or hydrolysates from marine fish and shrimps. Addition of these compounds to fish feed improves the food intake normally occurring when fish are given feed containing antibiotics. This observation may lead to possible improvements in aquaculture that are both economically and environmentally compatible. Thus, the occurrence of many biologically active peptides in marine byproduct hydrolysates is now well established, but numerous scientific and technological issues have to be resolved before these substances can be optimally exploited for human or animal nutrition and health. Biologically active substances or peptides from marine origin could be produced on an industrial scale as a consequence of the studies conducted on milk-derived proteins. These compounds could find applications both as dietary supplements in “functional foods” and as drugs, like bioactive peptides derived from milk proteins. However, the successful application of these bioactive peptides will require the demonstration of in vivo beneficial effects in animal models, so that the bioactivity in humans may be validated and potential adverse effects clarified. 4.2 Marine Waste as a Nutrient Source in Fermentation Processes Industrial fermentation processes require the availability of abundant and inexpensive substrate sources. As mentioned by Vecht-Lifshitz et al. [125], growth substrates constitute a major cost in the production of microbial cells and bioproductions; the nitrogen source tending to be the most expensive medium constituent. At present, commercial nitrogen sources are obtained from vegetable extracts, casein and slaughterhouse waste. Up to now, fish

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peptones have been investigated only to a minor extent, and their use in industrial processes is still poor despite their cheap price, as they constitute a waste product from the fish industry. However, the results of several studies have shown that in most cases fish peptones compared favourably to commercially available peptones produced from meat, casein or plant proteins [125–129]. The production of protease by Bacillus subtilis was strongly induced when cells were grown in media containing a non-defatted flour prepared from Sardinelle heads and viscera, compared to the same defatted fish meal or commercial peptones [130]. A shrimp waste acid hydrolysate containing 80% of glucosamine has been used as a carbon and energy source for the growth of Saccharomyces cerevisiae [131]. 4.3 Other Applications of FPHs Mackerel hydrolysates can be used to stimulate epoxy-pseudoisoeugenol(2-methylbutyrate) (EPB) production in transformed anise root culture [132]. EPB is potentially a valuable phenolic metabolite for regulation of nutraceutical-type phytochemicals during seed germination. Such phenolic metabolites are being targeted for food applications as antioxidants or as modulators of seed germination, to obtain seed-based functional phenolics for nutraceutical applications.

5 Genetic Traceability of Fish and Shellfish Species and By-Products Genetic identification methods provide new tools for an accurate and efficient determination of marine species specimens and also of derived products and coproducts (for a review, see [133]). As far as the characterisation of populations or the evaluation of stocks of commercially exploitable fishes are concerned, however, the use of genetic markers can lead to results that are often difficult to interpret and, therefore, could sometimes increase confusion more than solve difficulties. The tools are still new and one needs to establish consensus methods that have proved to be efficient and appropriate at each level of identification. Here, we aim to provide an understanding of a simple and reliable methodology using molecular genetics methods for the identification of marine species and derived processed products. Chemical signature and protein polymorphism have been already studied using denaturing electrophoresis [134, 135], isoelectric focusing [136] and high pressure liquid chromatography [137, 138]. These methods are powerful and acute, giving clues on the specimen history (chemical or biological environments impregnate metabolism or modify gene expression in organisms), but these parameters are too variable and too sensitive to be

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helpful for an efficient and easy identification at the level of population, species or upper [139]. On the other hand, molecules of nucleic acids are very stable, and a large number of markers can be found as ubiquitous sequences throughout living organisms. The accumulation of informative characters leads to preference for the comparison of sequences for species identification rather than use of methods showing a single nucleotide polymorphism (SNP)[140], or showing a fragment length polymorphism. The polymorphisms can be shown directly, as in the case of the characterisation of short tandemly repeated fragments (STR), of microsatellite markers [141–145], or by the characterisation of fragments issued by restriction digestion of a defined DNA fragment (RFLP) [146, 147]. Nevertheless, these methods appear to be well adapted for population structure analyses [144,148]. Methods based on protein characterisation have been also developed with the aim of identifying fish processed products obtained after heat denaturation [149] or cooking [150, 151]. In the case of fish product identification, these methods compete with DNA-based methods applied to smoked or canned samples [152–157]. 5.1 Choice of Marker Sequences Markers can be chosen in DNA extracted either from the nucleus or from mitochondria. Nuclear DNA is highly complex. Even if it can provide considerable systematic and phylogenetic information using the rhodopsin gene or the emergent use of RAG1 gene [158], its utilisation rather difficult for various reasons: its great variability among organisms, the fact that a high number of genes are unique in the genome and therefore difficult to isolate or amplify for further sequencing, and the fact that numerous genes exist as multi-gene families with several similar but different sequences. Some genes such as nuclear ribosomal RNA genes may occur inside one organism from several to a thousand copies without significant variability and can be found in all living materials. They are therefore good candidates for identification markers. Mitochondrial DNA occurs in the mitochondria of all eukaryotic cells. It is a simple molecule being a small circular DNA fragment typically containing 16 000 to 20 000 base pairs in length, haploid and maternally inherited. The mitochondrial genome contains only 37 genes, one non-coding region (the D-loop or control region), and is generally conserved and yet more variable than nuclear genes. It is generally considered to be a useful phylogenetic marker [159]. As mitochondria are numerous in each cell, the copy number is very high for each of these genes and, thus, the material is abundant and easy to extract, even from degraded (or processed) samples. The presence of protein-coding genes, rRNA-coding genes and non-coding fragments offers markers that are under different expression regulatory systems and therefore under different selective pressure mechanisms.

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Analysis of mtDNA, using RFLP analysis or DNA sequencing has indicated that some fragments contained in this genome may be more variable than others (Fig. 3). Genes encoding amino acid transfer RNAs (tRNA) are highly conserved among different species for their location as well as for their sequences. Mitochondrial ribosomal RNA genes show more variability: selective pressure acts only on these genes for the necessary conservation of the ribosomal RNA structure and allows changes in nucleotides for a high number of positions in the sequence. Structural genes coding for enzymes (the mitochondria contains the genes implicated in the process of oxidative phosphorylation, i.e. the production of energy and its storage) are more variable because most of the possible changes of the third nucleotide of several codons do not affect the translation meaning. The control region of the mitochondrial DNA (also named D-loop) contains DNA portions that are highly variable [160]. However, works performed on different fish species, i.e. Salmo trutta [161], Salmo salar [162] and Anguilla anguilla [163] have shown less variability in the non-coding D-loop than elsewhere in the mtDNA genome. This observation could be used in two ways. Firstly, one can concentrate on “slow evolving” parts of the genome in order to establish species comparisons or use the “fast evolving” portions for investigating populations. Secondly, and technically speaking, these types of structures are very helpful when searching parts of DNA to design PCR primers usable on a large variety of organisms. Starting from the weakly variable sequences of two transfer RNA genes flanking a more variable region of the mitochondrial genome, renders it easy to determine the sequences of this region from a large number of species, using the same couple of primers. Here, these primers could be tagged as “universal” primers as they could be used on different biological samples targeting the same genetic marker.

Fig. 3 Relative position of the genes largely used as genetic markers along the mitochondrial genome. Schematic representation of the mean nucleotide variability for the different genes

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The mitochondrial genomes of several commercial fish have now been completely sequenced, e.g. Atlantic cod [164], sea lamprey [165], salmon [166], sardine [167], shark (gummy shark [168], dogfish [169], spiny dogfish [170]) and various cartillagenous fish such as ray [171] and chimaera [172]. The portion represented in Fig.3 contains a set of genes, in particular cytochrome-b, largely used to establish phylogenetic relationship, determination of geographical population structure, genetic differentiation in fish, and for identification of species in nature and after processing [157, 160, 173–176]. Focusing on a ribosomal gene, i.e. 16SrDNA, a similar type of organisation with more variable–more stable regions is observed (Fig. 4). A portion of the 16SrDNA from 28 shark species representing three different orders was sequenced, aligned and compared. The analysis of the variability in nucleotide for each of the position along the sequence (indexed by a level of disorder) shows that parts of this gene are very stable (and could be used to design external PCR primers or internal sequencing primers necessary to achieve the determination of the sequence in case of long enough fragments) framing more variable regions whose changes in nucleotide are frequent and highly informative. An example of identification of an unknown “shark” processed product is presented below. Several studies on various specimens of this family revealed a low level of evolution of the mitochondrial genome [177] and the risk of using some nuclear genes for phylogeny [178]. The interest in using genetic markers for population studies [179–182] and for identification of species [183] or parts of morphologically similar specimens [184] has been demonstrated. In the domain of fish traceability, identification remains a basic problem strongly exacerbated when it comes to identifying detached shark fins or the finless and headless shark carcasses in fish markets. It is also a

Fig. 4 Schematic nucleotide variability for each of the different positions along the 16SrDNA fragment, expressed as entropy value using the 28 shark sequences selected from our shark sequence data library

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problem in the identification of the origin of manufactured and processed products with the aim of conservation and management controls on shark catch and trade. 5.2 Protocols for Fish and Fish Coproduct Identification With marine organisms, one frequently faces DNA extraction difficulties due to the presence in tissues of large quantities of mucus polysaccharides. An adapted protocol from the method described by Jones [185] using the non-ionic detergent CTAB (cetyltrimethylammonium bromide with formula C14 H29 N(CH3 )3 Br) proved to be very effective for these tissues. Starting with a pinpoint amount of muscle, skin, liver, bone or cartilaginous material, tissue was dissolved in 600 µL of buffered CTAB solution (2% (w/v) CTAB, 100 mM Tris-HCl, pH = 8, 20 mM EDTA, pH = 8 and 1.4 M NaCl), with addition of 0.2% 2-mercapto-ethanol. The sample was incubated for 30 min to 1 h at 60 ◦ C in the presence of 20 µL of a solution of 10 mg/mL of proteinase K. Protein extraction was carried out with two successive rounds of mixing an equal volume of chloroform/Isoamylol (24/1) and centrifugation (14 000 × g for 10 min). The upper aqueous solution was recovered and DNA precipitated with one tenth volume of NaOAc (3 M at pH = 5.2) and two volumes of ethanol. After homogenisation of the solution and centrifugation, the DNA was washed twice with 1 mL of 70% ethanol, dried and solubilised in 200 µL TE (Tris-HCl 10 mM, EDTA 10 Mm). The purity of the nucleic acid extract can be estimated by establishing a UV spectrum of the solution. DNA exhibits a maximum of absorption at 257 nm and ratios OD257/OD280 and OD257/OD230 should be the highest (between 1.8 and 2). The whole extraction protocol can be carried out within 1 or 2 h, depending on the difficulty of solubilisation of the tissues. This protocol is illustrated with data obtained on a processed (boiled, washed and dried) sample of “dry shark fin” collected on a fish market and used to test our method of identification of a marine processed product for which it is impossible to obtain any indication of origin (Fig. 5). In order to test the reliability of the extraction and homogeneity of the biological product, two different samples were taken and analysed separately. In order to obtain a large amount of a defined DNA fragment, a prior amplification of the selected marker was performed using PCR technology. The ready-to-go system (Amersham) was used where one has only to add DNA (1 µL), the two primers (2 µL) of a 10 nM solution) and water to a final volume of 25 µL. In the example shown here, a 22 mersens primer (5 AGGCAAGTCGTAACATGGTAAG3 ) whose target sequence is located in the 3 part of the 12 SrDNA and a 23 mer-antisens primer (5 ATCCAACATCGAGGTCGTAAACC3 ) with a target sequence in the 5 part of the 16 SrDNA were used. These sequences were determined according to

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Fig. 5 Dry fin sample. This product is found on the market, tagged as shark’s fin, imported from Thailand

the alignment of several mitochondrial sequences deposited in the gene bank. The expected DNA product should be around 1550 bp in length. The duration of such a program was around 3–4 h. Control of the result of a successful DNA amplification can be performed optionally by electrophoresis on a 1% agarose gel in TAE buffer [186]. After electrophoresis (1 h, 100 V), DNA fragments are stained with ethydium bromide (Fig. 6). The size of the fragments and an estimation of the amount of DNA produced can be determined by comparison with standard size markers (PstI digested Lambda phage DNA). When using a high (60 ◦ C) extension temperature, the amplification is highly specific and no interfering DNA fragments are observed. In the case of both samples, DNA extractions and PCR amplifications were successful. DNA can be easily extracted from the gel [187]. After excision of the DNA-containing agarose fragment with a scalpel, dissolution in 600 µL of a solubilisation mixture (QIAquick Gel Extraction Kit from Qiagen), and addition of 200 µL of isopropanol, the solution was loaded on an adsorption column. After a centrifugation (15 s), the fixed DNA was washed twice with a saline/ethanol solution, dried and eluted with a small volume (80 µL) of pure water or TE buffer. This DNA can be directly sequenced using either of the external primers that were used for the amplification process or using an internal primer. The sequencing here was achieved on a ABI PRISM 310 genetic analyser (Applied Biosystem) using a long read sequencing-61 cm capillary. Sequencing reactions were performed with PCR technology using the purified DNA fragment, one primer and a mixture of deoxynucleotides and fluorescent specifically labelled dideoxynucleotides, with a PCR program: 94 ◦ C × 30 s, 50 ◦ C × 30 s, 60 ◦ C × 4 min and 40 cycles. After precipitation, the reaction products were washed, dried, taken in 15 µL of formamide and subjected to electrophoresis

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Fig. 6 Agarose gel electrophoresis of the 16SrDNA fragment obtained from the dry fin sample after PCR amplification, EtBr staining and UV transillumination. The 1550 bp fragment is characterized using size standard fragments generated after Pst I digestion of Lambda phage DNA. Two different samples were set apart from the dry fin product to test heterogeneity

in the genetic analyser. The result was obtained as a processed electrophoregram (Fig. 7a). After checking for possible misinterpretations of the results of the electrophoresis, the sequences obtained in an easily exploitable form (Fig. 7b) from the two dry fin samples were aligned and compared. The first point to note is evidence of the real validity of the protocol used for extraction, amplification and sequencing for a variety of fresh tissues and also for ethanol precipitated, dehydrated and even boiled samples. For the dry fin samples, there were two different sequences for the two samples taken from the product. Between the two sequences, over 520 aligned nucleotides (data not shown) and 36 informative positions (2 deletions and 34 changes) are observed. Gaps in the alignment, as non-existing nucleotide in a particular sequence, can only be validated when aligning several sequences. Once fully valid as confirmed with an insured alignment, the sequences can be used as informative characters as defined changes of nucleotide. As a result, the two sequences obtained from this manufactured product appeared markedly different with 6.9% of change on the extent of the sequenced fragment. To determine the origin of the product, one can compare the sequence obtained with all the sequences stored in the data library. The search for homology could be carried out in seconds using an internal program such as BLAST [188]. Still very recently, the result of this research would have given the table presented in Fig. 8.

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Fig. 7 (a) Electrophoretic results of the direct sequencing of a portion of the 16SrDNA fragment after PCR amplification using our “shark universal” primers S1 and R4 (b) Corresponding dry fin 1 sequence under an edited format

The result of the search placed, with the best scores, the four shark sequences present in the data library, then the only available sequence from a rajidae, then the numerous sequences from bony fish. This search indicated that our unknown sample sequence came from a shark and that this shark could be a Carcharhiniforme. However, as only three orders (of the eight existing) are represented in the data library, it could arise from another order not present. A rapid browse of the content of the library shows that more than 100 000 sequences are from fish. Among them, more than 3000 contained at least a part of the 16 SrRNA gene, and close to 200 are sequences of the

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Fig. 8 Fourteen best scores of homology obtained after the search with “Blast n” using the unidentified dry fin 16SrDNA sequence 1 against the gene bank “other vertebrates”

complete mitochondrial genome. Some 867 sequences concern fish D-loop, more than 600 are part or complete cytochrome-b sequences. Nuclear genes such as RAG-1 (20) or rhodopsin (1678) genes are also represented in different ways. Even with a large number of sequences, bony fish (27 000 species) and cartilaginous fish (1000) species are only partially represented and the search gave a very inaccurate result. Moreover, the order seen for this output of the “Blast” search could be strongly biased because the comparison of the “personal sequence” with the data library sequences could be done using sequences covering different parts of the given marker (slow- or fast-evolving sequences) or fragment of different length. Another point is the alignment of some of the sequences with homologous (supposedly from the same species) sequences from the data library. Differences are sometimes observed that could only be explained by inaccurate sequencing analysis or inappropriate manipulation or errors in species identification. To eliminate these risks of error, it is thus advisable to ensure the identification of the specimens and to preserve the specimens used for the genetic signature determination in collection. Using the same program (Blast) we searched the sequence library that had been established for phylogenetic purposes [189]. In this case, using a set of sequences restricted to sequences from sharks that are supposed to be used to produce shark fins, the search placed first the sequences from Sphyrnidae, then numerous sequences from Carcharhinidae (Fig. 9). The very

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Fig. 9 Result of the homology search with the program “Blast n” using the unidentified dry shark fin sequence 2 against our MBS (Marine Biology Station) library. All the reference sequences used are with the same number of nucleotides. The score is very high for the comparison with the Sphyrna zygaena reference sequence because the whole dry fin sequence 2 is perfectly homologous to this one

high score obtained with the Sphyrna zygaena sequence is due to a perfect alignment of this sequence with that of the processed product. In the case of the other sample obtained from this product, it appeared that it also came from a Sphyrnidae, but not one represented in our sequence library. The result could be clearly shown by establishing a sequence similarity matrix expressing (as percentages) the levels of similitude of the different sequences (Fig. 10). From a rapid view of the percentages of similarities between the different sequences given by the sequence similitude matrix, it is possible to estimate interspecific variability at different levels, i.e. genus, family and order. From this comparison of shark sequences, using only one (partial) marker sequence, a level of nucleotidic changes of around 16.6% is observed when comparing Orectolobiformes to Carcharhinoformes and around 19.1% when comparing Orectolobiformes to Lamniformes. At the level of the order, the comparison of sequences from two representatives is between 16 and 19%. Inside an order (Lamniformes for example), 7.8 to 8% of changes are observed between the four studied families. At the level of the species, comparing the 11 species from the genus Carcharhiniformes, one observed a mean value of 4.8% of changes between these sequences. Highest values are obtained when

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Fig. 10 Similarity matrix of the sequences (expressed in percent). Cplu Carcarhinus plumbeus, Nacu Negaprion acutidens, Fin1 dry fin sample 1, Szyg Sphyrna zygaena, Fin2 dry fin sample 2, Cobs Carcarhinus obscurus, Cgal Carcarhinus galapensis, Tobe Triaenodon obesus, Ofer Odontaspis ferox, Ctau Carcharias taurus, Avul Alopias vulpinus, Cmax Cetorhinus maximus, Ipau Isurus paucus, Lnas Lamna nasus, Rtyp Rhincodon typus, Slew Sphyrna lewini, Csor Carcarhinus sorrah, Gcuv Galeocerdo cuvier

comparing sequences for other genus (7.3% for Sphyrna, 7.1% for Alopias and 6.7% for Isurus). These values for interspecies variability are very close to those for interfamily variability. This could be due to the fact that these families evolved a long time ago and a large number of the species are now extinct. In contrast, Carcharhinus has a large number of living species and a short evolution history (divergence time: 144 My). Applied to identification of the processed sample, the similarity matrix immediately shows a 100% similarity between the dry fin 2 sample and the Sphyrna zygaena sequence. This is evidence for the presence of fin product from this species in the unknown sample. The best observed score of similitude concerning the dry fin 1 sample is found with the Sphyrna lewini sequence (95.2%), then with the other Carcharhiniformes, then with the Lamniformes, in accordance with morphological characters based phylogeny. This shows that the processed sample is also composed from a second hammershark from the genus Sphyrna, probably Sphyrna mokaran, another species largely used for preparation of shark fin product in Asia. A 100% similitude is also observed between two sequences of reference (Carcharinus obscurus and Carcharhinus galapensis). As another genetic marker (Cytochrome-b gene) gave the same result for these two close species, we should come back to the collected specimen to ensure that no error could have occurred during the sampling of tissues. If not, complementary experiments could lead to a decision of synonymy.

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In conclusion, the use of genetic markers for species identification is a powerful, reliable and easy technique on fresh, ethanol-preserved, dry, boiled or even processed samples. It is, however, necessary to draw up consensual protocols and to adapt the choice of markers to the level of the need for identification. In addition, it is of primary importance that the sequences used as reference are given with all the precision wanted on specimens perfectly identified and preserved indefinitely in collection.

6 Conclusion In this review, we have attempted to survey the rapidly developing area of seafood by-product upgrading. Chemical processes have been applied with limited success for the recovery of seafood wastes and the production of quality products from them. Because of this, it is expected that biological processes will be prominent among those which will have to be developed for seafood waste processing. The general development of biotechnologies and the great diversity of commercial proteases should accelerate the application of biotechnological processes to fisheries. As far as species identification and traceabilty of fish and fish by-products are concerned, several European programs are dealing with the use of genetic markers for studying genetic diversity and the population structure of marine resources for stock identification and fisheries management. All these programmes are focused on single model species (trout, lobster, horse mackerel, redfish, cod or herring). Only one (Fishtrace) aims to establish a publicly accessible database compiling cytochrome-b and rhodopsin sequences from 150 fish species for identification purposes. There is a need to complete this approach, assessing methodologies for acquiring data on a large panel of commercial fish and shellfish species and related by-products. We hope that the above survey adequately demonstrates the potential applications of marine waste molecules in the field of nutrition and flavour. In addition to these well-established uses, we have shown that a significant research effort is currently being made to demonstrate the huge existing scope for the applications of biologically active peptides or molecules in the food, pharmaceutical and nutraceutical industries. However, the difficulties typically associated with supplying constant quality by-products remain the major practical obstacles for the introduction of these new products to the market place. A further major difficulty will be the development of economic processes to produce high added value hydrolysates in a repeatable manner. Currently, the processes employed are still at a laboratory scale, and therefore most of the final products purified or semi-purified are still not available as commercial products. It can be expected that in the next few years, bioactive peptide production and purification will be scaled

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up in collaboration with some companies who are convinced that it is absolutely necessary to upgrade the entire fish or shrimp caught (i.e. from fillets to viscera, including heads and frames). The growing demand for significant quantities of peptides of natural origin in the developed countries is encouraging these biotechnological alternatives. Thus, the economy of industrial fish and shrimp processing will be improved by the full utilisation of wastes and of underutilised marine organisms. Acknowledgements This work was performed within the integrated research project SeafoodPlus, Contract No. FOOD-CT-2004-506359. The partial financing of this work by the European Union is gratefully acknowledged. In addition, we thank Mr. Jean-Jacques Le Yeuc’h for reviewing the English language of this document. The identification and traceability programme “IDTRAMER” is a “research programme of regional interest of the Brittany region, France”.

References 1. Taylor T, Alasalvar C (2002) In: Alasalvar C, Taylor T (eds) Seafoods – quality, technology and nutraceutical applications. Springer, Berlin Hedelberg New York, p 123 2. Venugopal V (1997) Trends Food Sci Technol 8:271 3. FAO Annual Report (2002) The state of world fisheries and aquaculture (SOFIA), World review of fisheries and aquaculture, part 1, p 51 4. Shahidi F (1995) Protein concentrates from underutilized aquatic species In: Charalambous G (ed) Food flavors: generation, analysis and process influence. Elsevier, Amsterdam, p 1441 5. Sikorski ZE, Naczk M (1981) CRC Crit Rev Food Sci Nutr 14:201 6. Raa J, Gildberg A (1982) CRC Crit Rev Food Sci Nutri April:383 7. Venugopal V, Shahidi F (1995) CRC Crit Reviews Food Sci Nutr 35:431 8. Haard NF (1998) Food Technol 52:64 9. Simpson BK, Haard NF (1984) J Appl Biochem 6:135 10. Simpson BK, Haard NF (1984) Can J Biochem Cell Biol 62:894 11. Gildberg A (1992) Bioresource Technol 39:271 12. Arunchalam K, Haard NF (1985) Comp Biochem Physiol 80B:467 13. Guérard F, Le Gal Y (1987) Comp Biochem Physiol 88:823 14. Guérard F, Le Gal Y (1989) Biochimie 7:767 15. Hajjou M, Smine A, Guérard F, Le Gal Y (1995) Comp Biochem Physiol 110:791 16. Haard NF, Dimes LE, Arndt RE, Dong FM (1996) Comp Biochem Physiol 115B:533 17. Kristinsson HG, Rasco BA (2000) J Food Biochem 24:177 18. Smine A, Guérard F, Le Gal Y (1993) J Mar Biotechnol 1:41–46 19. Simpson BK, Haard NF (1987) In: Knorr D (ed) Food biotechnology. Marcel Dekker, New-York p 495 20. Gelman A, Mokadi S, Cogan U (1989) Comp Biochem Physiol 94B:113 21. Pavlisko A, Rial A, Coppes Z (1997) J Food Chem 21:383 22. Hennessey JR, Siebenaller JF (1987) J Exp Zoology 241:9 23. Squires EJ, Haard NF, Feltham LAW (1986) Can J Biochem Cell Biol 64:210 24. Myrnes B, Johansen A (1994) Prep Biochem 24:69 25. Vilhelmesson O (1997) Trends Food Sci Technol 8:266

Fish and Shellfish Upgrading, Traceability

159

26. Haard NF Simpson BK (1994) In: Martin AM (ed) Fisheries processing – biotechnological applications. Chapman & Hall, p 132 27. Petersen BR, The impact of the enzymic hydrolysis process on recovery and use of proteins. pp 149-175 28. Mackie IM (1982) Process Biochem Jan/Feb: 26 29. Kristinsson HG, Rasco BA (2000) CRC Crit Rev Food Sci Nutr 40:43 30. Shahidi F, Synowiecki J (1996) Food Chem 57:317 31. Vieira GHF, Martin AM, Saker-Sampaiao S, Omar S, Goncalves RCF (1995) J Sci Food Agric 69:61 32. Martin AM, Porter D (1995) In: Charalambous G (ed) Food flavors: generation, analysis and process influence. Elsevier, Amsterdam, p 1395 33. Quaglia GB, Orban E (1987) J Sci Food Agric 38:263 34. Diniz FM, Martin AM (1998) Food Sci Technol Intern 4:91 35. Guérard F, Dufossé L, De La Broise D, Binet A (2001) J Mol Cat B: Enz 11:1051 36. Cassia RO, Martone CB, Sanchez JJ (2000) Latin Am Appl Res 30:241 37. Synowiecki J, Al-Khateeb NAAQ (2000) Food Chem 68:147 38. Shahidi F (1995) Extraction of value-added components from shellfish processing discards. In: Charalambous G (ed) Food flavors: generation, analysis and process influence. Elsevier, Amsterdam, p 1427 39. Gilberg A, Stenberg S (2001) Process Biochem 36:809 40. Vijayalakshmi MA, Lemieux L, Amiot J (1986) J Liq Chromatogr 9:3559 41. Pianelli G (1999) Agro-Food-Industry Hi-Tech – sept/oct:46 42. Diniz FM, Martin AM (1997) Agro-Food-Industry Hi-Tech – May/June:9 43. Adler-Nissen J (1986) Enzymatic hydrolysis of food proteins. Elsevier , New York, p 13 44. Hoyle NT, Merrit JH (1994) J Food Sci 59:76 45. Vieira GHF, Martin AM, Saker-Sampaiao S, Sobreira-Rocha CA, Goncalves RCF (1995) In: Charalambous G (ed) Food flavors: generation, analysis and process influence. Elsevier, Amsterdam, p 1405 46. Shahidi F, Han XQ, Synowiecki J (1995) Food Chem 53:285 47. Liaset B, Lied E, Espe M (2000) J Sci Food Agric 80:581 48. Kristinsson HG, Rasco BA (2000) J Agric Food Chem 48:657 49. Guérard F, Ravallec-Plé R, De La Broise D, Binet A, Dufosse L (2001a) In: Thonart P, Hofman M (eds) Engineering and manufacturing for biotechnology, focus on biotechnology, vol IV. Kluwer, p 39 50. Benjakul S, Morrissey MT (1997) J Agric Food Chem 45:3423 51. Diniz FM, Martin AM (1996) Int J Food Sci Technol 31:419 52. Ravallec-Plé R, Charlot C, Pires C, Braga V, Batista I, Van Wormhoudt A, Le Gal Y, Fouchereau-Péron M (2001) J Sci Food Agric 81:1120 53. Shahidi F, Synowiecki J, Balejko J (1994) J Agric Food Chem 42:2634 54. Guérard F, Guimas L, Binet A (2002) J Mol Cat B: Enz 19-20:489 55. Liaset B, Nortvedt R, Lied E, Espe M (2002) Process Biochem 37:1263 56. Simpson BK, Nayeri G, Yaylayan V, Ashie INA (1998) Food Chem 61:131 57. Baek HH, Cadwallader KR (1995) J Food Sci 60:929 58. Diniz FM, Martin AM (1997) Int J Food Sci Nutr 48:191 59. Adler-Nissen J (1982) J Chem Tech Biotechnol 32:138 60. Panyam D, Kilara A (1996) Trends in Food Sci Technol 7:120 61. Adler-Nissen J (1979) J Agric Food Chem 27:1256 62. Nielsen PM, Petersen D, Dambmann C (2001) J Food Sci 66:642 63. Silvestre MPC (1997) Food Chem 60:263

160

F. Guérard et al.

64. Fujinari EM, Manes JD (1995) In: Charalambous G (ed) Food flavors: generation, analysis and process influence. Elsevier, Amsterdam, p 929 65. Shahidi F, Naczk M, Pegg RB, Synowiecki J (1991) Food Chem 42:145 66. FitzGerald RJ (1998) Int Dairy J 8:451 67. Meisel H (1993) In: Sawatzki G, Renner B (eds), New perspectives in infant nutrition. Thieme, Stuttgart, p 153 68. Pihlanto-Leppälä A (2001) Trends in Food Sci Technol 11:347 69. Yamamoto N (1997) Inc Biopoly 43:129 70. Yamamoto N, Ejiri M, Mizuno S (2003) Curr Pharm Design 9:1345 71. Meisel H (1997) Livestock Prod Sci 50:125 72. Meisel H (2001) Austral J Dairy Technol 56:83 73. Clare DA, Swaisgood HE (2000) J Dairy Sci 83:1187 74. Clare DA, Catignani GL, Swaisgood HE (2003) Curr Pharm Design 9:1239 75. Floris R, Recio I, Berklout B, Visser S (2003) Curr Pharm Design 9:1257 76. Schlimme E, Meisel H (1995) Nahrung Food 39:1 77. Bordenave S, Fruitier I, Ballandier I, Sannier F, Gildberg A, Batista I, Piot JM (2002) Prep Biochem Biotechnol 32:65 78. Dorman T, Bernard L, Glaze P, Hogan J, Skinner R, Nelson D, Bowker L, Head D (1995) J Adv Med 8:193 79. Le Poncin M (1996a) Eur Neuropsychopharmacol 6:110 80. Le Poncin M (1996b) Eur Neuropsychopharmacol 6:187 81. Bernet F, Montel V, Noel B, Dupouy JP (2000) Psychopharmacology 149:34 82. Maruyama S, Mitachi H, Awaya J, Suzuki H (1987) Agric Biol Chem 51:2557 83. Meisel H, FitzGerald RJ (2003) Curr Pharm Design 9:1289 84. Kohmura M, Nio N, Ariyoshi N (1990) Agric Biol Chem 54:835 85. Miyoshi S, Ishikawa H, Kaneko T, Fukui F, Tanaka H, Mayurama S (1991) Agric Biol Chem 55:1313 86. Matsumara N, Fujii M, Takeda Y, Sugita K, Shimizu T (1993a) Biosci Biotech Biochem 57:695 87. Kohama Y, Oka H, Kayamori Y, Tsujikawa, Mimura T Nagase Y, Satake M (1991) Agric Biol Chem 55:2169 88. Matsui T, Matsufuji H, Seki E, Osajima K, Nakashima M, Osajima Y (1993) Biosci Biotech Biochem 57:922 89. Matsumara N, Fujii M, Takeda Y, Sugita K, Shimizu T (1993b) Biosci Biotech Biochem 57:1743 90. Wako Y, Ishikawa Q, Muramoto K (1996) Biosci Biotech Biochem 60:1353 91. Jeon YJ, Byun HG, Kim SK (2000) Process Biochem 35:471 92. Hata Y, Yamamoto M, Ohni M, Nakajima K, Nakamura Y, Takano T (1996) Am J Clin Nutr 64:767 93. Seppo L, Kerojoki O, Suomalainen T, Korpela R (2002) Milchwissenschaft 57:124 94. Kawasaki T, Seki E, Osajima K, Yoshida M, Asada K , Matsui T (2000) J Human Hypertens 14:519 95. Kayser H, Meisel H (1996) FEBS Letter 383:18 96. Coste M, Rochet V, Léonil J, Mollé D, Bouhallab S, Tomé D (1992) Immunol Lett 33:41 97. Gildberg A, Bøgwald J, Johansen A, Stenberg E (1996) Comp Biochem Physiol B: Biochem Mol Biol 114:97 98. Bogwald J, Dalmo R, McQueen Leifson R, Stenberg E, Gildberg A (1996) Fish Shellfish Immunol 6:3 99. Franco-Cereceda A, Gennari C, Nami R, Agnusdei D, Pernow J, Lundberg JM, Fischer JA (1987) Circ Res 60:393

Fish and Shellfish Upgrading, Traceability 100. 101. 102. 103. 104. 105. 106. 107. 108. 109. 110. 111. 112. 113. 114. 115. 116. 117. 118. 119. 120. 121.

122. 123. 124. 125. 126. 127. 128. 129. 130. 131. 132. 133. 134. 135.

161

Hughes JJ, Levine AS, Morley JE, Gosnell B, Silvis SE (1984) Peptides 5:665 Umeda Y, Arizawa M (1989) Jap J Pharmacol 51:377 Nong YH, Titus RG, Ribeiro JMC, Remold HG (1989) J Immunol 143:45 Roos BA, Fischer JA, Pignat W, Alander CB, Raisz LG (1986) Endocrinol 118:46 Fouchereau-Péron M, Duvail L, Michel C, Gildberg A, Batista I, Le Gal Y (1999) Biotechnol Appl Biochem 29:87 Rousseau M, Batista I, Le Gal Y, Fouchereau-Péron M (2001) Electronic J Biotechnol (EJB) 4:1 Cancre I, Ravallec R, Van-Vormhoudt A, Stenberg E, Gildberg A, Le Gal Y (1999) Mar Biotechnol 1:489 Ravallec-Plé R, Gilmartin L, Van Wormhoudt A, Le Gal Y (2000) J Sci Food Agric 80:2176 Park PJ, Jung WK, Nam KS, Shahidi F, Kim SK (2001) JAOCS 78:651 Chen HM, Muramoto K, Yamauchi F, Nokihara K (1996) J Agric Food Chem 44:2619 Chen HM, Muramoto K, Yamauchi F, Fujimoto K, Nokihara K (1998) J Agric Food Chem 46:49 Okada Y, Okada M (1998) J Agric Food Chem 46:401 Rival SG, Boeriu C, Wichers HJ (2001) J Agric Food Chem 49:295 Chen HM, Muramoto K, Yamauchi F(1995) J Agric Food Chem 43:574 Chuang WL, Pan B S, Tsai JS (2000) J Food Biochem 24:333 Shahidi F, Amarowicz R (1996) JAOCS 73:1197 Amarowicz R, Shahidi F (1997) Food Chem 58:355 Kim SK, Kim YT, Byun HG, Nam KS, Joo DS, Shahidi (2001) J Agric Food Chem 49:1984 Li SJ, Seymour TA, King AJ, Morrissey MT (1998) J Food Sci 63:438 Seymour TA, Li SJ, Morrissey M (1996) J Agric Food Chem 44:682 Suetsuna K (2000) Mar Biotechnol 2:5 Guérard F, Sumaya-Martinez MT, Binet A (2002) In: Colliec-Joualt S (ed) Marine biotechnology: an overview of leading fields. Actes du Colloque Européen IXth ESMB, Marine Biotechnology, Nantes, France, 12-14 May 2002. IFREMER, INIST 0761-3962, p 155 Yoshikawa T, Naito Y, Masui K, Fujii T, Boku Y, Nakagawa S, Yoshida N, Kondo M (1997) Biomed Pharmacother 51:328 Pokorny J (1991) Trends Food Sci Technol 2:223 Guérard F, Sumaya-Martinez MT (2003) JAOCS 80:467 Vecht-Lifshitz, Almas KA, Zomer E (1990) Lett Appl Microbiol 10:183 Gilberg A, Batista I, Strom E (1989) Biotechnol Appl Biochem 11:413 Dufossé L, De La Broise D, Guérard F (1997) Recent Res Develop Microbiol 1:365 Dufossé L, De La Broise D, Guérard F (2001) Current Microbiol 42:32 De La Broise D, Dauer G, Gildberg A, Guérard F (1998) J Mar Biotechnol 6:111 Ellouz Y, Bayoudh A, Kammoun S, Gharsallah N, Nasri M (2001) Bioresource Technol 80:49 Ferrer J, Paez G, Marmol Z, Ramones E, Garcia H, Forster CF (1996) Bioresource Technol 57:55 Andarwulan N, Shetty K (2000) Food Biotechnol 14:1 Hastein T, Hill BJ, Berthe F, Lightner DV (2001) Rev Sci Tech 20:564 Verrez-Bagnis V, Ladrat C, Morzel M, Noel J, Fleurence J (2001) Electrophoresis 22:1539 Etienne M, Jérome M, Fleurence J, Rehbein H, Kundiger R, Malmhedem-Yman I, Ferm M, Craig A, Mackie I, Jessen F, Smelt A, Luten J (1999) Electrophoresis 20:1923

162

F. Guérard et al.

136. Lunstrom R (1980) J Assoc Off Anal Chem 63:69 137. Osman MA, Ashoor SH, Marsh PC (1987) J Assoc Off Anal Chem 70:618 138. Pineiro C, Sotelo CG, Medina I, Gallardo JM, Pérez-Martin R (1997) Z Lebensm Unters Forsch A 204:411 139. Rehbein H, Etienne M, Jérome M, Hattula T, Knudsen B, Jessen F, Luten JB, Bouquet W, Mackie IM, Ritchie AH, Martin R, Mendes R (1995) Food Chem 52:1193 140. Rehbein H, Kress G, Schmidt T (1997) J Sci Food Agric 74:35 141. Yu H-T, Lee Y-T, Huang S-W, Chiu T-S (2002) Mar Biotechnol 4 471 142. Yue GH, Li Y, Chao TM, Chou R, Orban L (2002) Mar Biotechnol 4:503 143. Rivera MAJ, Graham GC, Roderick GK (2003) Mar Biotechnol 5:126 144. Sekino M, Hamaguchi M, Aranishi F, Okoshi K (2003) Mar Biotechnol 5:227 145. Delghandi SR, Mortensen A, Westgaard J-I (2003) 5:141 146. Lopez-Pinon MJ, Insua A, Mendez J (2002) Mar Biotech 4, 495-502 147. Klinbunga S, Khamnamtong N, Tassanakajon A, Puanglarp N, Jarayabhand P, Yoosukh W (2003) Mar Biotechnol 5:27 148. Was A, Wenne R (2003) Mar Biotechnol 5:234 149. Pineiro C, Barros-Velazquez J, Pérez-Martin R, Martinez I, Jacobsen T, Rehbein H, Kundiger R, Mendes R, Etienne M, Jérome M, Craig A, Mackie I, Jessen, F (1999) Electrophoresis 20:1425 150. Rehbein H, Kundiger R, Malmhedem-Yman I, Ferm M, Etienne M, Jérome M, Craig A, Mackie I, Jessen F, Martinez I, Mendes R, Smelt A, Luten J, Pineiro C, PérezMartin R (1999) Food Chem 67:333 151. Etienne M, Jérome M, Fleurence J, Rehbein H, Kundiger R, Mendes R, Costa H, Pérez-Martin R, Pineiro-Gonzales C, Craig A, Mackie I, Malmhedem-Yman I, Ferm M, Martinez I, Jessen F, Smelt A, Luten J (2000) J Agric Food Chem 48:653 152. Etienne M, Jérome M, Fleurence J, Rehbein H, Kundiger R, Mendes R, Costa H, Martinez I (2001) Food Chem 72:105 153. Lockley AK, Bardsley RG (2000) Trends Food Sci Technol 11:67 154. Mackie I, Craig A, Etienne M, Jérome M, Fleurence J, Jessen F, Smelt A, Kruijt A, Malmheden-Yman I, Fern M, Martinez I, Pérez-Martin R, Pineiro C, Rehbein H, Kundiger R (2000) Food Chem 71:1 155. Quintero J, Sotelo CG, Rehbein H, Pryde SE, Medine I, Pérez-Martin R, Rey-Méndez M, Vidal R, Mackie IM (1998) J Agric Food Chem 46:1662 156. Ram JL, Ram ML, Baidoun F (1996) J Agric Food Chem 44:2460 157. Jerome M, Lemaire C, Verrez-Bagnis V, Etienne M (2003) J Agric Food Chem 25:7326 158. Bernstein RM, Schulter SF, Bernstein H, Marchalonis J (1996) Proc Natl Acad Sci 93:9454 159. Zardoya R, Meyer A, (1996) Mol Biol Evol 13:933 160. Murgia R, Tola G, Archer SN, Vallerga S, Hirano J (2002) Mar Biotechnol 4:119 161. Hall HJ, Nawrocki LW (1995) J Fish Biol 46(2):360 162. O’Connel M, Danzmann RG, Cornuet JM, Wright JM, Ferguson MM (1997) Can J Fish Aquat Sci 54:1391 163. Daemen E, Cross T, Ollevier F, Volckaert F (2001) Mar Biol 139:755 164. Johansen S, Bakke I (1996) Mol Mar Biol Biotechnol 5:203 165. Lee WJ, Kocher TD (1995) Genetics 139:873 166. Hurst CD, Bartlett SE, Davidson W, Bruce IJ (1999) Gene 239:237 167. Inoue JG, Miya M, Tsukamoto K, Nishida M (2000) Fisheries Sci 66:924 168. Cao Y, Waddell PJ, Norihiro O, Hasegawa M (1998) Mol Biol Evol 15:1637 169. Delarbre C, Spruyt N, Delmarre C, Gallut C, Barriel V, Janvier P, Laudet V, Gachelin G (1998) Genetics 150:331

Fish and Shellfish Upgrading, Traceability 170. 171. 172. 173. 174.

175. 176. 177. 178. 179. 180. 181. 182. 183. 184. 185. 186. 187. 188. 189.

163

Rasmussen AS, Arnason U (1999) J Mol Evol 48:118 Rasmussen AS, Arnason U (1999) Proc Natl Acad Sci USA 96:2177 Arnason U, Gullberg A, Janke A (2001) Zool Scr 30:249 Barlett SE, Davidson WS (1991) Can JF Aqua Sci 48:309 Sotelo CG, Calo-Mata P, Chapela MJ, Perez-Martin RI, Rehbein H, Hold GL, Russell VJ, Pryde S, Quinteiro J, Izquierdo M, Rey-Mendez M, Rosa C, Santos AT (2001) J Agric Food Chem 49:4562 Sebastio P, Zanelli P, Neri TM (2001) J Agric Food Chem 49:1194 Sanjuan A, Comesana AS (2002) J Food Prot 6:1016 Martin AP, Naylor GJP, Palumbi SR (1992) Nature 357:153 Martin AP, Burg TM (2002) Syst Biol 51:570 Heist EJ, Musick JA, Graves JE (1996) Fishery Bull 94:664 Heist EJ, Musick JA, Graves JE (1996) Can J Fish Aquat Sci, 53:583 Gardner MG, Ward RD (1998) Mar Freshwater Res 49:733 Heist EJ, Gold JR (1999) Copeia 1999:182 Martin AP (1993) NOAA Technical Report NMFS 115:53 Pank M, Stanhope M, Natanson L, Kohler N, Shivji M (2001) Mar Biotechnol 3:231 Jones AS (1953) Biochim Biophys Acta 10:607 Southern E (1975) J Mol Biol 98:503 Smith HO (1980) Methods Enzymol 65:371 Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ (1990) J Mol Biol 215:403 Iglesias SP, Lecointre G, Sellos DY (2005) Mol Phyl Evol 34(3):569

Adv Biochem Engin/Biotechnol (2005) 96: 165–188 DOI 10.1007/b135784 © Springer-Verlag Berlin Heidelberg 2005 Published online: 25 August 2005

Marine Microalgae Tadashi Matsunaga1 (u) · Haruko Takeyama1 · Hideki Miyashita2 · Hiroko Yokouchi1 1 Department

of Biotechnology, Tokyo University of Agriculture and Technology, Koganei, 184-8588 Tokyo, Japan [email protected]

2 Department

of Technology and Ecology, Hall of Global Environmental Studies, Kyoto University, Yoshida-Honmachi, Sakyo-ku, 606-8501 Kyoto, Japan

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Production of Useful Chemicals by Marine Cyanobacteria . . . . . . . . . . . . . . . . Rhodophyta . . . . . . . . . . . . . . . . . Chlorophyta . . . . . . . . . . . . . . . . . Cryptophyta . . . . . . . . . . . . . . . . . Heterokontophyta . . . . . . . . . . . . . . Dinophyta . . . . . . . . . . . . . . . . . . Haptophyta . . . . . . . . . . . . . . . . . Euglenophyta . . . . . . . . . . . . . . . .

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Metabolic Engineering of Marine Microalgae Gene Transfer Methods for Marine Microalgae Metabolic Engineering of Marine Microalgae for Producing Valuable Metabolites . . . . . . Whole genome analyses in marine microalgae

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Microalgal Mass Cultivation Technologies . . . . . . . . . . . . . . . . . . Open Culture Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Photobioreactor (Closed Culture Systems) . . . . . . . . . . . . . . . . . .

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CO2 Fixation Using Microalgal Cultures in Industry . . . . . . . . . . . .

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Abstract Marine microalgae, the largest primary biomass, have been attracting attention as resources for new metabolites and biotechnologically useful genes. The diversified marine environment harbors a large variety of microalgae. In this paper, the biotechnological aspects and fundamental characteristics of marine microalgae are reviewed. Keywords Marine microalgae · Useful material production · Genetic manipulation · Mass cultivation · Photoreactor

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1 Introduction The primary producers of oxygen in aquatic environments are algae, especially planktonic microalgae. These microorganisms are widely distributed in nature and have adapted to different environments with great diversity in size, morphology, life cycle, pigments, and metabolism. About one half of global photosynthesis and oxygen production is accomplished by marine microalgae. They play an important role in CO2 recycling through photosynthesis, which is similar to higher plants in O2 -evolved systems (PSI and PSII). Research in microalgae has been carried out not only on physiological aspects but also to develop production of useful biomaterials. The advantages of their utilization in production are (1) their ability to convert CO2 to useful materials through photosynthesis and (2) their ability to grow in natural environments under inorganic conditions. For example, marine microalgae can be cultivated using seawater, CO2 , and sunlight. Recent developments in the biotechnology of microalgae have been focused on their production of useful materials applicable to the cosmetic and medical fields. Genetic modification and molecular tools have been developed mainly in eubacterial microalgae, cyanobacteria (blue-green algae). In contrast, genetic modification has been only gradually applied to eukaryotic microalgae. Recently, whole genome sequences and EST analyses have been performed in marine strains. The elucidation of genomewide information may help in the development of new biotechnological applications using microalgae. In this chapter, we review the useful applications of microalgae for genetic engineering, cultivation technologies, and CO2 fixation as follows: 1. 2. 3. 4.

Production of useful chemicals by marine microalgae Metabolic engineering of marine microalgae Microalgal mass cultivation technologies CO2 fixation using microalgal cultures in industry

2 Production of Useful Chemicals by Marine Microalgae 2.1 Cyanobacteria The cyanobacteria are oxygenic photosynthetic prokaryotes that show large diversity in their morphology, physiology, ecology, biochemistry, and other characteristics. Currently, more than 2000 species are recognized, which comprise two fifths of known bacterial species (5000 species). Such variation complicates estimation of species diversity. Cyanobacteria contain chlorophyll a,

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phycobiliproteins, and characteristic glycosylated xanthophylls, such as myxoxanthophyll and oscillaxanthin [1–3]. The phycobiliproteins are watersoluble pigments consisting of red-colored phycoerythrin and blue-colored phycocyanin and allophycocyanin. They have a characteristic structural feature, the phycobilisome, that is used as a photosynthetic light-harvesting antenna. Cyanobacteria are unicellular, multicellular, colonial, and branched or unbranched filamentous forms. Specialized cells, heterocystous and akinates, are contained in some of the filamentous-form cells. Some cells produce extracellular matrix such as sheaths, capsules, and slimes that consist mainly of polysaccharides. Cyanobacteria are distributed widely not only in salt water but also freshwater, brackish water, polar regions, hot springs, and deserts. Some also exist as symbionts in sponges, ascidians, echiuroid worms, planktonic diatoms, and dinoflagellates in marine environments [4] and lichens and azollae in terrestrial environments. Cyanobacteria, especially marine pelagic Synechococcus and Prochlorococcus, contribute largely to global oxygen production. Many commercial applications have been proposed for marine cyanobacteria, although no marine strain presently is commercially supplied. Recent proposals to evaluate potential commercial uses typically fall into three categories: bioactive chemical compounds [5–10], polysaccharides [11–14], and evaluation of new genes for recombinant expression. Cyanobacteria can produce a large variety of complex chemical compounds. Bioactive compounds isolated from marine cyanobacteria have recently been summarized by Burja et al. [5] and Takeyama and Matsunaga [15]. Novel plant growth regulators that promote redifferentiation, germination, and plantlet formation [16], tyrosinase inhibitors [17], UV-A absorbing compounds [18], sulfated polysaccharides showing anti-HIV activity, and novel antibiotics with light-regulated activity [19] are among the many compounds that have been studied. Matsunaga et al. [20] reported that several marine cyanobacterial strains, such as Phormidium sp. NKBG 041105 and Oscillatoria sp. NKBG 091600, showed high cis-palmitoleic acid content (54.5% and 54.4% of total fatty acid, respectively). The discovery of biochemically active compounds from marine cyanobacteria, including enzyme inhibitors, herbicides, antimycotics, antifeedants, multi-drug-resistance reversers, and antimalarial and immunosupressive agents, has dramatically increased over the last few years. This has been due to the adaptation and use of current cyanobacterial collections and cyanobacteria-derived compounds for screening in new pharmaceutical and industrial assays. The focus on the polysaccharides of marine cyanobacteria as well as freshwater strains also has greatly increased in relation to interest in their exploitation for various industrial applications [11–14]. Cyanobacteria produce three types of extracellular matrix consisting mainly of polysaccharides, which have unique bio- and physicochemical characteristics. Most of them are com-

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posed of at least ten different monosaccharides and contain pentoses, which have not been observed in other prokaryotic polysaccharides. The anionic nature of these unique polysaccharides is due to the presence of acidic sugars and anionic organic and inorganic compounds. Little work has been devoted to potential applications of marine cyanobacterial polysaccharides. Extracellular polysaccharide production by cyanobacteria, as well as their possible applications, was reviewed by Philippis and Vincenzini [11] and Philippis et al. [13]. Marine cyanobacteria also are attractive as a resource for useful enzymes and genes [21–29]. Cyanobacteria commonly produce complex macromolecules that often possess biological activities such as cytotoxicity or microbial toxicity [5]. Recent genetic analyses have revealed that most of these macromolecules, as well as unusual small molecules, are coded for by large gene clusters representing nonribosomal peptide synthetase and polyketide synthetase [21–23, 25, 26, 28]. These gene clusters may be manipulated for the production of new chemicals. Many varieties of the gene clusters are present in most cyanobacteria [21, 24]. Therefore, cyanobacteria are attractive not only as producers of useful bioactive macromolecules and enzymes but also for production of complex macromolecules that may become important pharmaceuticals. 2.2 Rhodophyta The rhodophytes, or red algae, contain chlorophyll a, carotenoids, and phycobiliproteins. Their red color is due to the presence of phycoerythrin in the outermost part of the phycobilisomes, while other regions of the algae look blue-green due to the absence of phycoerythrin. Rhodophytes are unicellular, or composed of simple or complex filamentous aggregates. Flagellated cells have not been observed. Red macroalgae commonly inhabit tropical and temperate zones near shores. About 600 genera and 5500 species have been recognized. Most of them (98%) are marine macroalgal species. Red macroalgae are of economic significance [30]. Porphyra and a few other species are cultured for human food. The production of red algal polysaccharides such as agar, agarose, and carrageenans is also an important industry. These compounds are widely used for laboratory cell culture media, nucleic acid purification, or food processing, respectively. Although rhodophyte microalgae have not been produced commercially as yet, their polysaccharides are considered to have commercial potential [31–35]. For example, in concentrated solutions of polysaccharides from the unicellular rhodophyte Porphyridium sp., viscosity is stable over a wide range of pH, temperature, and salinity. These properties indicate possible applications for use as a thickening agent in aqueous systems or as a stabilizer for emulsions and suspensions [31]. In addition to its potential application as

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a viscosity stabilizer, the polysaccharide and the biomass of Porphyridium sp. have been used as functional food additives [32]. The colon and jejunum changed morphologically with hypertrophy in the muscularis layer in rats fed diets containing pelleted biomass or sulfated polysaccharides. In addition, it was shown in rats that the algal polysaccharide and biomass were potent hypocholesterolemic agents active at low concentrations in the diet. Moreover, the sulfated polysaccharide of Porphyridium sp. has shown promising antiviral activity against a variety of animal viruses including Herpes simplex viruses types 1 and 2 (HSV 1, 2) Varicella zoster virus (VZV), and HIV types 1 and 2. The compounds also showed significant inhibition of productive infection with retroviruses (murine leukenia virus, HIV-1, and HIV-2) and cell transformation by murine sarcoma virus in vitro [33–35]. Thus, red microalgae and their polysaccharides seem to be good candidates for the development of antiviral drugs. 2.3 Chlorophyta Cells of chlorophytes are green due to chlorophyll a and b, the same predominant photosynthetic pigments as those of land plants. Some algae show yellowish-green or red-green colors due to the presence of a certain amount of carotenoids such as β-carotene, prasinoxanthin, siphonaxanthin, and astaxanthin. These chlorophyta form starch in the chloroplast as a storage product of photosynthesis. Chlorophytes are unicellular, multicellular, colonial, filamentous, siphonous, and thallus. The Chlorophyta consist of five classes, the Prasinophyceae, the Ulvophyceae, the Chlorophyceae, the Trebouxiophyceae, and the Charophyceae. The Treboxiophyceae recently were separated from the Chlorophyceae. The Chlorophyta are primarily freshwater algae with approximately 500 genera comprising 16 000 species. Only about 10% of these are marine species. The Ulvophyceae are primarily multicellular marine green algae. In addition, some species from the Prasinophyceae, Chlorophyceae, and Trebouxiophyceae families are found in the marine environment. Depending on the class, it has been estimated that there would be at least two to three times more species in this phylum. A marine species of the Chlorophyceae, Dunaliella, has been cultivated commercially for food supplements and β-carotene production [36]. The microalgal biomass of some marine Tetraselmis and Pyramimonas strains in the Prasinophyceae family also are used for fish food additives [37]. Recently, anti-inflammatory and immunosupressive properties were discovered in the extracts [38] or extracted polysaccharides [39] of another marine species, Chlorella stigmatophora. Miura et al. [40] reported that Chlorella sp. NKG 042401 contains 10% γ -linolenic acid (C18 : 3), which is present in the cells mainly in the form of galactolipid.

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2.4 Cryptophyta Cryptophytes are unicellular, cryptomonad flagellates with 12 to 23 genera comprised of 200 species. A few species are colorless heterotrophs, but most possess various colored plastids with chlorophylls a and c, carotenoids, and phycobiliprotein. Alloxanthin is a xanthophyll that is unique to cryptomonads. Morphologically, cells have a flattened asymmetrical shape with two anterior flagella, slightly unequal in length. They are distributed widely both in freshwater and marine environments. Approximately half of the known species inhabit marine environments. It has been estimated that there would be about six times more species in this phylum. Only a few strains such as Rhodomonas minuta and Cryptomonas sp. have been used for aquaculture feeds since they contain significant amounts of polyunsaturated fatty acids (PUFAs) [37]. No further application has been proposed. 2.5 Heterokontophyta The phylum Heterokontophyta is the most diverse algal group with huge commercial and biotechnological potentials [30, 37]. They range in size from microscopic unicells to giant kelp averaging several meters. Cells of heterokontophytes contain chlorophyll a with chlorophyll c and carotenoids such as fucoxanthin or vaucheriaxanthin. They are characterized primarily by the similarities in their ultrastructural and biochemical characteristics. Heterokontophyte microalgae are widely used as feed in mariculture/ aquaculture [30, 37, 41]. Diatoms such as Chaetoceros calcitrans, Chaetoceros gracilis, Chaetoceros muelleri, Skeletonema costatum, and Thalasiosira pseudodonana are commonly used as live feeds for all growth stages of bivalve molluscs (e.g., oysters, scallops, clams, and mussels), for crustacean larvae, and for zooplankton used as feed for larvae. The genera Navicula, Nitzschia, Cocconeis, and Amphora also are used to feed juvenile abalone. Some Eustigmatophyceae species of the genus Nannochloropsis are commonly fed to Artemia or rotifers, which in turn are fed to crustacean and fish larvae. The biotechnological potential of diatoms is also concerned with PUFA production. Most diatoms have a high content of eicosapentenoic acid (EPA) 20:5 (n-3). Phaeodactylum tricornutum and Nitzschia laevis especially have been investigated for EPA production. In addition, EPA production by diatoms has been reviewed recently by Lebeau and Robert [42, 43]. Recent advances in heterotrophic production of EPA by microalgae were also reviewed by Wen and Chen [44]. The Pinguiophyceae also have significant biotechnological potential for use in fish feed and for PUFA production [45]. Pinguiophyceae consist of five ma-

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rine unicellular algal species, Pinguiochrysis pyriformis, Phaeomonas parva, Pinguiococcus pyrenoidosus, Glossomastix chrysoplasta, and Polypodochrysis teissieri. These algae have an unusually high percentage of PUFAs, especially EPA. The EPA content ranges from 23.5% to 56.0% of the total fatty acids in these five species. They also contain arachidonic acid (AA) 20:4 (n-6) and docoahexaenoic acid (DHA) 22:6 (n-3). Pinguiococcus pyrenoidosus, Glossomastix chrysoplasta, and Polypodochrysis teissieri have docosatetraenoic acid (DTA) 22:4 (n-3) ranging from 4.4% to 9.5% of total fatty acid content. This significant oil balance, as well as their lack of cell wall, indicates that they are good candidates as food/feed sources. Evaluations of the utility of Pinguiophyceae for food/feed supplements and optimization of growth conditions necessary for efficient production are being carried out. Some marine colorless heterokontophytes also are being tested for the production of DHA [36, 46–48]. Schizochytrium, Thraustochytrium, and Ulkenia are representatives of the class Labyrinthulea. They produce substantial amounts of PUFAs, especially DHA [46–48]. The safety of biomass production and differences in oil content are intensively studied [49–53]. Utilization of fermented “okara” for DHA and/or EPA production by thraustochitrids also has been reported, while the yield was lower by growth in a glucoseyeast-extract medium than by fermentation [54]. 2.6 Dinophyta The Dinophyta include the dinoflagellates, most of which are unicellular, with two dissimilar flagella [1–3]. Flagellated cells show characteristic forward-spiraling swimming motions. Organisms in this phylum have remarkable morphological diversity including nonflagellate amoeboid, coccoid, palmelloid, or filamentous. Approximately 130 genera with about 2000 living and 2000 fossil species have been described in this group. Most are marine and only about 220 species are from freshwater. About half of the known species are colorless heterotrophs. Most of the plastid-containing phototrophic dinoflagellates contain chlorophylls a and c2 and carotenoids such as β-carotene and peridinin, the unique accessory xanthophyll in this phylum. Dinoflagellates are also characterized by cell coverings consisting of a layer containing many closely adjacent, flattened amphiesmal (thecal) vesicles. In many species, each amphiesmal vesicle contains a thecal plate composed of cellulose. About 60 dinoflagellate species are known to produce cytolytic, hepatotoxic, or neurotoxic compounds. Some form harmful red tides. The majority of toxin-producing dinoflagellates are photosynthetic, estuarine, or coastal shallow-water forms that are capable of producing benthic resting cysts and that tend to form monospecific populations. Freshwater species are not known to produce toxins. The dinophytes comprise autotrophs, mixotrophs, osmotrophs, phagotrophs, and parasites. Dinoflag-

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ellate endosymbionts known as zooxanthellae are essential for the existence of coral reef ecosystems. Symbiotic as well as parasitic forms also are present within the cells or tissues of fish, invertebrates, and filamentous algae. Biotechnological applications for dinoflagellates have not been intensively performed, probably because most dinoflagellates can not be easily cultured. The only exception is the DHA produced by heterotrophically grown Crypthecodinium cohnii [36]. DHA produced by these cells is distributed widely as food supplements, such as for infant formula. Recently, significant biological activities have been attributed to the polysaccharide from Gymnodinium sp. [55–58]. For example, strong cytotoxicity against several human leukemic cell lines leading to apoptosis [55] and potent anticancer activity mediated by the inhibition of topoisomerase I and II [58]. Optimal growth conditions for Gyrodinium impudicum were reported to produce a sulfated polysaccharide that showed antiviral activity against encephalomyocarditis virus [59]. 2.7 Haptophyta The phylum Haptophyta (haptophytes or prymnesiophytes) is a group of unicellular flagellates characterized by the presence of a haptonema between two smooth flagella [1–3]. The role of haptonema is unclear, although it sometimes functions as a feed-capturing net, in avoidance reaction by coiling and recoiling, and as an attachment organ on surfaces. There also are haptonemaless flagellates or nonflagellate unicells or colonies. The cells of haptophytes are brownish or yellowish-green containing chlorophylls a and c1 /c2 and carotenoids such as β-carotene, fucoxanthin, diadinoxanthin, and diatoxanthin. The cells are commonly covered with scales made mainly by carbohydrates or calcium bicarbonate. Many species known as coccolithophorids produce calcified scales called coccoliths. About 70 genera with 300 species have been recognized to date. Most are primarily marine species inhabiting tropical seawater. The group is distributed worldwide and is often an important source of food for aquatic communities. Some haptophytes, however, produce algal blooms and cause serious problems for fish and for fishermen by producing dimethyl sulfide (DMS), a noxious-smelling compound that affects fish migrations and alters their normal routes. Microalgal biomass of haptophytes is commonly used as living feed in aquaculture [37]. Isochrysis galbana and Pavlova lutheri, especially, are used as living feeds for bivalve molluscs, crustacean larvae and zooplanktons that in turn are used for crustacean and fish larvae. Some cells can produce PUFAs such as DHA or EPA. In addition, the DHA content in I. galbana was shown to be enhanced by low temperature or incubation of the culture in the dark after reaching plateau phase growth [60]. Furthermore, it was shown that these algae are useful for DHA enrichment of feed such as rotifers for the larvae of several marine fish species [61].

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2.8 Euglenophyta Euglenophytes are unicellular organisms with two pantonematic flagella arising from the bottom of a flask-shaped invagination called a “gullet.” A few have stages with colonies or are enclosed within a mucilaginous capsule. There are about 40 genera comprised of 900 species, of which one third have green plastids. The chloroplast originating from green algae contains chlorophylls a and b and carotenoids such as diadinoxanthin, neoxanthin, and β-carotene. Two thirds of the genera are heterotrophic, some having colorless plastids and some lacking plastids altogether, living either saprotrophically or phagocytically. Cell walls are absent, but there is a characteristic cell-covering structure called a pellicle, composed of protein-rich spiral strips beneath the cell membrane. One to several flagella may be present, and nonflagellate cells can undergo a type of motion involving changes in cell shape. Although most species are found in highly eutrophic freshwater environments such as ponds and ditches, some euglenoids have important ecological roles in particular marine environments. Very few euglenoids have been grown in axenic culture, and euglenoid culture media are generally very nutrient rich. No direct economic significance has been associated with this phylum, probably because of the difficulties in culturing. Although euglenoids generally are harmless, toxin production has been demonstrated in some freshwater Euglena sp. [62]. On the other hand, E. gracilis Z is one of the few microorganisms that simultaneously produces antioxidant vitamins such as β-carotene and vitamins C and E. Efficient production of these vitamins consists of a two-step culture under photoheterotrophic/photoautotrophic conditions [63].

3 Metabolic Engineering of Marine Microalgae 3.1 Gene Transfer Methods for Marine Microalgae Genetic studies on microalgae have been redirected mainly toward analysis of photosynthesis and metabolic pathways. A limited number of microalgae such as cyanobacteria have been used in biotechnological applications. Development of molecular techniques for physiological analysis and enhancement of biotechnological applications is necessary. Many attempts at gene transfer have been made in eukaryotic and prokaryotic microalgae. Genetic manipulation in prokaryotic microalgae cyanobacteria have been studied extensively after several transformable unicellular

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strains were discovered. At first, the freshwater cyanobacterium Synechococcus PCC7942 was reported to have an ability to take up DNA. Subsequently, several other naturally transformable freshwater strains have been found. Gene transfer has been developed mainly in freshwater strains, Synechococcus, Synechocystis, Anabaeba, and Nostoc [64]. Only a few marine cyanobacterial strains of the genus Synechococcus have been used for heterogeneous gene expression and other genetic applications [65, 66]. There are two commonly used gene transfer procedures: transformation using naturally occurring or artificially competent cells, e.g., conjugation with Escherichia coli, or physical methods for gene introduction, e.g., electroporation and particle bombardment. In marine cyanobacteria, natural transformation has been reported for Synechococcus sp. PCC7002 [67]. Other strains have been transformed successfully by electroporation or conjugation. Further, plasmids harvested from several marine microalgal species have been used as vector DNA for gene transfer. Many cyanobacteria-harboring endogenous plasmids have been reported. Some functional genes were found to be coded on freshwater cyanobacterial plasmids [68]; however, most cyanobacterial plasmids are cryptic. Marine plasmids have been found in Synechococcus sp. NKBG042902, which has high phycocyanin content and a rapid growth rate. Extracts from this strain promote plant germination [16]. This strain contains more than three cryptic endogenous plasmids, pSY09 (> 10 kbp), pSY10 (2.6 kbp), and pSY11 (2.3 kbp). Plasmid pSY10 has the unique replication characteristic that their copy number increases under high salinity conditions [70]. To investigate the function and replication mechanism of these plasmids, the plasmids pAQ1 (4.8 kbp) of the marine strain Synechococcus PCC7002 and pSY10 of NKBG 042902 were entirely sequenced [71, 73], and a gene transfer system using pSY10 was established [69]. The complete sequence of Synechococcus pSY10 (2561 bp) includes seven potential open reading frames (ORFs). The longest ORF has homology with the replication region of plasmids from several bacteria [72]. pSY10 did not hybridize with other plasmids purified from Synechococcus sp. NKBG 042902. Replication of these plasmids appears to be controlled by different mechanisms. Plasmids are maintained at high copy number in cyanobacteria, suggesting the possibility that they act as a shuttle vector between cyanobacteria and E. coli. In fact, a shuttle vector with E. coli has been constructed using pSY10. Conjugative gene transfer has been reported mainly for the freshwater filamentous cyanobacterium Anabaena PCC7120, which is not a naturally transformable strain. Conjugation in Anabaena PCC7120 was carried out using conjugal plasmids such as RP4 (IncP), a helper plasmid carrying a mobilization gene, and a shuttle vector carrying cyanobacterial replicons [74]. The other filamentous freshwater cyanobacteria Plectonema boryanum [75] and Fremyella diplosiphon [76] also can be successfully transformed by conju-

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gation. A conjugative plasmid vector in unicellular freshwater cyanobacteria has been constructed [77]. Conjugative gene transfer using a broad-host range vector pKT230 (IncQ, Kmr , and Smr ) was successful for the marine cyanobacterium Synechococcus sp. NKBG 15041C [78]. It was demonstrated that this plasmid is stably maintained in cyanobacterial cells [79]. This introduced a new tool in cyanobacteria biotechnology since most transformations were carried out using the shuttle vector plasmid containing a cyanobacterial plasmid origin of replication. Random gene insertion into the genome using the transposon vector pSUP 1021 (which carries the RP4-specific mob site) and the transposon Tn5 was demonstrated in NKBG 15041C. In marine cyanobacteria, besides the plasmid vector system, the construction of a phage vector system also is required to enable the cloning of large DNA fragments in specific cyanobacterial hosts. Since cyanophages were first reported by Safferman and Morris [80], various types of cyanophages have been found in seawater [81, 82] and have been characterized as to their genetic diversity and phylogenetic affiliations [83]. These vectors could be utilized for gene transfer in the near future. The particle gun or microprojectile method has been developed for delivering DNA into plant cells and tissues. In prokaryotes, Shark et al. [84] reported the biolistic transformation of Bacillus megaterium. Both gold and tungsten particles have been used as DNA carriers in this system. However, small particles are required for prokaryotic transformation. DNA (pSUP1021) conjugated onto nano-sized bio-magnetic beads (50–100 nm in diameter), purified from the magnetic bacterium Magnetospirillum sp. AMB-1 [85], was used to successfully transform a marine cyanobacterium, Synechococcus sp. NKBG15041c [86]. In eukaryotic microalgae, some unicellular and multicellular algae have been successfully transformed, although most of them are freshwater strains of diatom and chlorella. Marine strains, such as diatom Phaeodactylum tricornutum, Thalassiosira weissflogii, and green algae Dunaliella salina, have been reported to be transformed as well [87–90]. Particle bombardment and electroporation are used mostly for these microalgae. Stable transformation for the purpose of enhancing the production of useful materials or analyzing gene expression has been carried out. However, the level of protein production by transformants varied due to multiple insertion of the target gene into the genome and to variation in transcriptional efficiency caused by random integration.

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3.2 Metabolic Engineering of Marine Microalgae for Producing Valuable Metabolites Enhanced production of valuable primary or secondary metabolites in microalgae can be rendered possible by high density cultivation and/or application of genetic manipulation. Recent pharmaceutical interest in unsaturated fatty acids has triggered the search for sources of these valuable compounds. Several eukaryotic microalgae are known to produce highly unsaturated fatty acids such as EPA and DHA, which are valuable dietary components [44, 61]. Genetic engineering has been applied to produce EPA in the marine cyanobacterium Synechococcus sp. [66]. Cyanobacteria do not have the biosynthetic pathway to produce them. The EPA synthesis gene cluster (ca. 38 kbp) isolated from a marine bacterium Shewanella putrefaciens SCRC-2738 was cloned to the marine cyanobacterium using a broad-host cosmid vector, pJRD215 (10.2 kbp, Smr Kmr ). The cyanobacterial transconjugants grown at 29 ◦ C produced EPA only at 0.12 mg/g dry cell, whereas those grown at 23 ◦ C produced EPA at 0.56 mg/g dry cell. The content of EPA grown at 23 ◦ C increased to 0.64 mg/g dry cell after 24 h incubation at 17 ◦ C. Furthermore, EPA production was improved by partial deletion of the EPA gene cluster to stabilize its expression and maintenance in host cyanobacterial cells [91]. Most diatoms do not have the capacity to grow on exogenous glucose in the absence of light. The transformable marine diatom Phaeodactylum tricornutum exhibited heterotrophic growth after the introduction of a single gene for glucose transporters glut1 or hup1 [92]. For this purpose, plasmid (pPha-T1; glut1-gfp) was introduced into P. tricornutum by using a biolistic procedure, and transformants were selected for zeocin resistance in the light. Exogenous glucose entering the transformants can be metabolized at a high rate of flux, allowing the cells to thrive in the absence of light. The trophic conversion of microalgae, such as diatoms, is a critical first step in engineering algae for successful large-scale cultivation. The genetic engineering of microalgae for industrial purposes also has been performed in freshwater cyanobacteria where the ketocarotenoid astaxanthine, an extremely efficient antioxidant, was synthesized by introduction of the beta-c-4-oxygenase gene (crtO) from the green alga Haematococcus [93]. Ethylene production also was demonstrated in the cyanobacterium Synechococcus elongates PCC7942 by chromosomal insertion of an ethyleneforming enzyme [94]. However, the reaction catalyzed by the ethyleneforming enzyme induced metabolic stress that was detrimental to the host cell.

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3.3 Whole genome analyses in marine microalgae Sequencing of microbial genomes has become a routine procedure for gene discovery. The most abundant population in marine cyanobacteria is Prochlorococcus, which are the smallest phytoplankton known, with a diameter of about 0.6 µm. Now the complete genomes of three strains of Prochlorococcus [95, 96] and one strain of Synechococcus [97] have been sequenced and analyzed. The information obtained from genome sequences and subsequently by comparative genome analyses takes on importance as the 2000 genes of these minimal life units are sufficient to generate the most abundant global biomass from solar energy and inorganic compounds. The genome database for cyanobacteria is available at (http://www.kazusa.or.jp/cyano/cyano.html). In eukaryotic microalgae genomics, the genome composition of the genetically transformable diatom strain Phaeodactylum tricornutum was analyzed by the generation of approximately 1000 express sequence tags (ESTs) [98]. Interestingly, many sequences were shown to have more similarity with animal genes than with their plant counterparts.

4 Microalgal Mass Cultivation Technologies Photosynthetic microorganisms play an important role in the conversion of solar energy into chemical energy. Photosynthetic conversion is an efficient and alternative process used in several industrial fields. Attempts to develop large-scale methodologies for the cultivation of microalgae have been performed using many different kinds of cultivation systems for providing alternatives to fermentation and agriculture products [99]. Algal biomass has historically served as fertilizer [100] and a food source for both humans and animals [101, 102] for secondary waste water treatment [103] and bioremediation [104, 105]. This use of algal biomass is an important consideration for industrial applications of microalgal cultures. With advances in processing technology, algal biomass has come to be seen as a possible source of fuels, fine chemicals, and pharmaceuticals [106]. Several species of microalgae, which produce useful chemicals such as amino acids, vitamins, carotenoids (β-carotene), fatty acids (DHA, EPA, γ -LA 18:3 (n-6)), polysaccharides, and antibiotics have been reported. Many microalgal products have already been commercialized [36, 107–109]. Further, microalgal production of energy resources has been extensively investigated. Development of processes that utilize the majority of the resulting microalgal biomass as energy sources would be preferred. Such processes may allow the recycling of evolved

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CO2 from human energy consumption rather than direct emission, as is the present case for fossil fuels. The following six products for use as fuels can be produced from microalgal biomass: hydrogen (through biophotolysis), methane (through anaerobic digestion), ethanol (through yeast or other alcohol fermentation), triglycerides (through extraction of lipids), methyl ester fuels (through transesterification of lipids), and liquid hydrocarbons (from Botryococcus braunii). The development of efficient culture systems is necessary for algal mass production and the industrial applications of microalgae. The growth rate and maximum biomass yield of microalgal strains are affected by culture parameters (light, temperature, and pH) and nutritional status (CO2 , nitrogen, and phosphate concentration). On the other hand, increasing the density of cultures decreases photon availability to individual cells. Light penetration of microalgal cultures is poor, especially at high cell densities, and such poor photon availability decreases specific growth rates. Higher biomass yields can be expected if sufficient photons are provided in high density cultures of microalgae. Large-scale culture systems have been constructed (classified as open and closed systems) with the greatest attention directed to the light supply (Fig. 1) [110]. Strains such as Chlorella, Scenedesmus, Dunaliella, Spirulina, Porphyridium, and Haematococcus have been cultured using photobioreactors to obtain several useful materials. 4.1 Open Culture Systems Several different types of open culture systems have been proposed (Fig. 1a-d). These open culture systems are the simplest method of algal cultivation and offer advantages in low construction cost and ease of operation [111]. The open culture systems require large surface areas and shallow depth (ca. 12–15 cm) to improve light penetration. Furthermore, agitation of the culture prevents the cells from sinking to the bottom and facilitates efficient cell growth with sunlight. The raceway pound has been developed into various types, where those employing a paddle wheel for agitation have been used most frequently for outdoor production of microalgae [112, 113]. The raceway pound for commercial production of microalgae requires an area of 1000 to 5000 m2 . Contamination by different algal species and other organisms is the biggest problem in open culture systems, and therefore Chlorella, Dunaliella, and Spirulina, which are tolerant to extreme conditions (high nutrient concentrations, high salinity, and high pH), are especially desirable strains for open culture systems. Vonshak et al. [114] demonstrated that contamination by Chlorella in outdoor Spirulina cultures was prevented by maintaining the culture medium at high bicarbonate concentration (0.2 M). Grazers sometimes

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Fig. 1 Illustration of algal mass culture systems

found contaminants in Spirulina cultures could be arrested by addition of ammonia (2 mM). The open culture system is easily affected by weather conditions. For example, rain dilutes salinity, causing contamination. Outdoor open culture systems are chosen mainly for production of food sources in aquaculture [43, 115, 116]. However, several algae producing useful chemicals require more restricted conditions for efficient growth and for metabolite production.

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4.2 Photobioreactor (Closed Culture Systems) Closed systems have been expected to overcome the disadvantages of open culture systems, and several types of photobioreactors have been devised. EPA-producing microalgae diatoms have been cultured at various scales in photobioreactors [43]. The closed system is required here because the species used have no selective advantages like those of Dunaliella and Spirulina. Figure 1e–i shows an example of large-scale closed photobioreactors. These photobioreactors offer several advantages: (1) facilitate maintenance of monoalgal cultures by protecting them from contamination; (2) reduce water loss and the subsequent increase of salinity in the culture medium; (3) result in higher productivity with greater cell densities, reducing harvesting costs; and (4) are applicable to various microalgal species under favorable culture conditions. However, for an efficient and reliable large-scale culture system, several criteria need to be considered [117] such as light utilization efficiency, homogeneous mixing (turbulence), low shear environment, temperature control, and efficient gas transfer. The production yield of algal biomass depends on the light path length to each cell, and therefore the surface-to-volume ratio is an important factor for efficient light utilization in photobioreactors. The productivity of photobioreactors is determined by the light regime inside the bioreactors. In addition to the light regime, oxygen accumulation and shear stress limit productivity in certain designs [118]. Tubular reactors have been refined, and the diameter of a tubular reactor is now less than 40 cm. Richmond et al. demonstrated that reduction in the tube diameter from 5.0 cm to 2.8 cm enhanced the biomass yield 1.8 times [119]. Narrower tube diameters may increase efficient light utilization as well as promote a faster flow rate, enhancing the algal productivity and reducing fouling on the inside wall of the tubes. In tubular reactors, flow rates of 30–50 cm/s generally are used and airlift is the most effective circulation method of the culture rather than using centrifugal, rotary positive displacement, and peristaltic pumps. The main advantages of airlift systems are their low shear and relative simplicity of construction. Several modifications in tube arrangement have been carried out for optimizing light penetration. Until recently, most of the tubular reactor was laid on the ground such that the lower part of the tubes received less light than the upper part. Torzillo et al. constructed a two-plane tubular bioreactor for optimizing light availability, where each tube in the lower plane is placed in the vacant space between two tubes in the upper plane [120]. They showed that this two-plane tubular bioreactor (145 L of culture volume) has an effective surface-to-volume ratio (49 L/m) resulting in a net volumetric productivity of 1.5 g dry wt/L/d using Spirulina platensis. The helical tubular reactor shown in Fig. 1e consists of a vertical tower coiled up within a lone tube, increasing the land use efficiency. When biomass

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productivity of various tubular reactors was compared on a footprint basis, the values were in the range of 15 to 30 g dry wt/m/d for all reactor types. The effect of temperature on biomass yield is significant in algal culture. The culture in the tubular reactor often is maintained at higher temperatures than that in the open raceway. Spirulina cultured in the tubular reactor could be warmed faster than in the open raceway up to 35–37 ◦ C, the optimal range for growth [119]. The closed reactors are sometimes overheated and thus are more suitable for the thermophilic or thermotolerant strains. Temperature control using a heat exchanger and/or evaporative cooling by spraying water onto the surface is sometimes required for the cultivation of general strains [120–122]. The effect of hydrodynamic stress on two different microalgae strains, Dunaliella tertiolecta and D. salina, also was investigated [121]. The data demonstrated that bubble rising and bubble bursting were not responsible for cell death. Regarding nozzle diameter, small nozzles were more detrimental to cells. Bubble formation at the sparger was the main cause of cell death. A problem in the closed system is photooxidative damage to the cells caused by accumulation of dissolved oxygen produced by photosynthesis during the light period. In the open system, evolved oxygen is diffused easily to the atmosphere. By contrast, because oxygen cannot escape from closed reactors, degasser systems sometimes are required. The culture part of closed photobioreactors has been constructed with several materials such as glass, methyl-polymethacrylate, polyethylene, polypropylene, vinyl-polychlorine, silicone, and stainless steel. These photobioreactors have been designed for optimal utilization of external illumination like sunlight. Optical fibers also have been employed as internal light sources. Photobioreactors employing optical fibers have the advantage of controlling illumination and light period duration and have a high surfacearea-to-volume ratio [123]. The efficiency of light utilization of microalgae also was studied under light/dark cycles encountered in airlift photobioreactors using D. tertiolecta [124]. Optimization of cultivation parameter growth kinetics has increased productivity in photobioreactors. The Accelerationstat (A-stat) cultivation method has been proposed to determine the culture steady state where the dilution rate is increased at a constant rate under light-limiting conditions in a photobioreactor [125]. Open pond systems have lower productivity of algal biomass, require larger land areas, and involve higher land costs. By contrast, closed culture systems can achieve high-density culture and the overall volume of algal culture can be reduced, resulting in decreased land costs. However, a certain amount of land area is still required for collecting a sufficient amount of solar energy, and therefore operating costs are higher than for open systems. Moreover, solar radiation, temperature, and other factors regulating algal productivity are significantly affected by location. Suitable culture systems

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should be chosen according to the target products and available environmental conditions.

5 CO2 Fixation Using Microalgal Cultures in Industry The possible use of biological CO2 fixation to reduce anthropogenic CO2 emission has been investigated. However, the questions related to CO2 reduction on the basis of global net amount are debated because biomass must be decomposed and CO2 is released into the atmosphere as a result. For development of onsite CO2 fixation systems using microalgae, efficient photobioreactors and strains that can fix large quantities of CO2 are required. Several applied studies have been conducted that consider the direct biological utilization of CO2 in emission gases from coal-fired power plants and the steel and cement plants that produce large quantities of CO2 , NOx , and SOx (inhibitory gases for photosynthesis). Therefore, microalgae that can grow under such extreme conditions will be required for direct CO2 fixation. Many microalgal strains capable of rapid growth in water sparged with emitted gases and under other extreme conditions such as high pH have been screened, and a marine alga Chlorococcum littorale showing high CO2 tolerance and high growth rate in the linear growth phase was obtained [126]. In the report of the IEA Greenhouse Gas R&D Program, a system for the reduction and recycling of CO2 emission from coal-fired power plants was designed, where CO2 fixed products generated by microalgal culture are used as biomass fuels, which will substitute eventually for fossil fuels [110]. Costs of microalgae CO2 mitigation using the designed systems have been estimated based on several design specifications such as (1) plant size, (2) gas condition, (3) conditions for CO2 biofixation, (4) plant operation, (5) CO2 production rate, and (6) algal strain (Nannochloropsis sp). Analyses of the designed system showed that CO2 mitigation costs closely depend on productivity of algae and solar radiation. In spite of these recent advances, microalgal strains that can achieve higher photosynthetic efficiency at higher solar radiation are necessary. In addition, a photobioreactor, in which microalgae converts CO2 to biomass at high photosynthetic efficiency, also is required. One of the applications for CaCO3 recycling also was demonstrated [127, 128]. Coccolithophorids are unicellular planktonic marine algae that produce elaborate structures called coccoliths comprised of scales or plates of CaCO3 . In the oceans, huge blooms of coccolithophorid algae occur that have been recognized as contributing to ocean floor sediment formation. Therefore, algae play an important role in the global carbon cycle by CaCO3 recycling. CO2 fixation by artificial weathering of waste concrete and coccolithophorid algae cultures has been proposed (Fig. 2) [129]. During artificial

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Fig. 2 Design of CO2 fixation by artificial weathering of waste concrete and culture of coccolithophorid algae

weathering of waste concrete suspended in seawater, atmospheric CO2 can be absorbed and dissolved as bicarbonate ions, which are a major source of coccolith particles. Coccolithophorid algae can use bicarbonate ions to form CaCO3 particles. Consequently, CO2 absorbed by artificial weathering can be mineralized and fixed permanently. Artificial weathering of waste concrete also is a useful method to supply bicarbonate ions to cells of the coccolithophorid alga Emiliania huxleyi. CO2 fixation by artificial weathering of waste concrete and coccolithophorid algae cultures can be applied to the reduction of CO2 emission from cement plants (Fig. 3). Coccoliths can be used as an alternative to limestone, which is a carbonate source used for cement production. In the cement industry, CO2 is produced mainly by decomposition of limestone during the burning of cement clinker. If CaCO3 recycling can be achieved by artificial weathering of waste concrete and coccolithophorid culture, CO2 emissions by the cement industry might be reduced. It has been estimated that the amount of CO2 absorbed by the weathering of waste concrete is greater than that of CO2 emitted during a cement production when CO2 reduction and recycle systems using microalgae are implemented. Moreover, CO2 is absorbed by the coccolithophorid alga cultures themselves [128]. Glucose oxidase and uricase have been immobilized onto purified ultrafine coccolith particles to illustrate their potential as a support material for biotechnological application [130]. If microalgal biomass can be stored in concrete without the decomposition of the biomass back to CO2 , removal of anthropogenic CO2 may be achieved. Extensive studies of biological CO2 fixation using microalgal cultures have been pursued. A primary goal is the complete removal of CO2 in discharged gas emitted by such an onsite system. Because of the land area requirements and a CO2 mitigation cost of $264 per ton as carbon, it is difficult at present to apply microalgal cultures for CO2 removal. Most nations are seriously concerned about the increase of atmospheric CO2 concentration, and intensive efforts to reduce the anthropogenic CO2 emissions are being made. Microalgae culture may be one of the important processes facilitating such efforts [131]. Increasing attention is being paid to resource sustainability in all

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Fig. 3 Design of CO2 fixation by artificial weathering of waste concrete and culture of coccolithophorid algae

industries, and developing new technologies for microalgal culture will help to provide sustainable resources.

References 1. Graham L, Wilcox L (2000) Algae. Prentice-Hall, Englewood Cliffs, NJ 2. Hoek C van den, Mann D, Jahns H (1995) Algae: An Introduction to Phycology. Cambridge University Press, Cambridge, UK 3. Lee R (1999) Phycology. Cambridge University Press, Cambridge, UK 4. Raven J (2002) Biol Environ 102B:3 5. Burja A, Banaigs B, Abou-Mansour E, Burgess J, Wright P (2001) Tetrahedron 57:9347 6. Skulberg O (2000) J Appl Phycol 12:341 7. Mundt S, Kreitlow S, Nowotny A, Effmert U (2001) Int J Hyg Environ Health 203:327 8. Proksch P, Edrada R, Ebel R (2002) Appl Microbiol Biotechnol 59:125 9. Han B, McPhail K, Ligresti A, Marozo V, Gerwick W (2003) J Nat Prod 66:1364 10. Volkman J (2003) Appl Microbiol Biotechnol 60:495 11. De Philippis R, Vincenzini M (1998) FEMS Microbiol Rev 22:151 12. Shah V, Ray A, Garg N, Madamwar D (2000) Curr Microbiol 40:274 13. De Philippis R, Sili C, Paperi R, Vincenzini M (2001) J Appl Phycol 13:293 14. Otero A, Vincenzini M (2003) J Biotechnol 102:143 15. Takeyama H, Matsunaga T (1998) Production of useful materials from marine microalgae. Vijay Primlani for Oxford & IBH Publishing, New Delhi 16. Wake H, Akasaka A, Umetsu H, Ozeki Y, Shimomura K, Matsunaga T (1992) Plant Cell Reports 11:62 17. Wachi Y, Sode K, Horikoshi K, Takeyama H, Matsunaga T (1995) Biotechnol Tech 9:633

Marine Microalgae

185

18. Wachi Y, Burgess J, Iwamoto K, Yamada N, Nakamura N, Matsunaga T (1995) Biochim Biophys Acta 1244:165 19. Matsunaga T, Sudo H, Takemasa H, Wachi Y, Nakamura N (1996) Appl Microbiol Biotechnol 45:24 20. Matsunaga T, Takeyama H, Miura Y, Yamazaki T, Furuya H, Sode K (1995) FEMS Microbiol Lett 133:137 21. Neilan B, Dittmann E, Rouhiainen L, Bass R, Schaub V, Sivonen K, Borner T (1999) J Bacteriol 181:4089 22. Berg H, Ziegler K, Piotukh K, Baier K, Lockau W, Volkmer-Engert R (2000) Eur J Biochem 267:5561 23. Doekel S, Marahiel M (2001) Metab Eng 3:64 24. Christiansen G, Dittmann E, Ordorika L, Rippka R, Herdman M, Borner T (2001) Arch Microbiol 176:452 25. Chang Z, Flatt P, Gerwick W, Nguyen V, Willis C, Sherman D (2002) Gene 296 26. Oppermann-Sanio F, Steinbuchel A (2002) Naturwissenschaften 89:11 27. Hutchinson C (2003) Proc Natl Acad Sci USA 100:3010 28. Hoffmann D, Hevel J, Moore R, Moore B (2003) Gene 311:171 29. Hook I, Ryan S, Sheridan H (2003) Phytochemistry 63:31 30. Wikfors G, Ohno M (2001) J Phycol 37:968 31. Arad S, Friedman O, Rotem A (1988) Appl Environ Microbiol 54:2411 32. Dvir I, Chayoth R, Sod-Moriah U, Shany S, Nyska A, Stark A, Madar Z, Arad S (2000) Br J Nutr 84:469 33. Huleihel M, Ishanu V, Tal J, Arad S (2001) J Appl Phycol 13:127 34. Talyshinsky M, Souprun Y, Huleihel M (2002) Cancer Cell Int 2:8 35. Huleihel M, Ishanu V, Tal J, Arad S (2002) J Biochem Biophys Methods 50:189 36. Apt K, Behrens P (1999) J Phycol 35:215 37. Brown MR (2002) Nutritional value of microalgae for aquaculture. In: Cruz-Suárez LE, Ricque-Marie D, Tapia-Salazar M, Gaxiola-Cortés MG, Simoes N (eds) Advances en Nutrición Acuicola VI. Memorias del VI Simposium Internacional de Nutrición Acuicola. 3–6 September 2002, Cancun, Quintana Roo, Mexico, p 282 38. Guzman S, Gato A, Calleja J (2001) Phytother Res 15:224 39. Guzman S, Gato A, Lamela M, Freire-Garabal M, Calleja J (2003) Phytother Res 17:665 40. Miura Y, Sode K, Nakamura N, Matsunaga N, Matsunaga T (1993) FEMS Microbiol Lett 107:163 41. Browitzka M (1997) J Appl Phycol 9:393 42. Lebeau T, Robert J (2003) Appl Microbiol Biotechnol 60:612 43. Lebeau T, Robert J (2003) Appl Microbiol Biotechnol 60:624 44. Wen Z, Chen F (2003) Biotechnol Adv 21:273 45. Kawachi M, Inouye I, Honda D, O’Kelly C, Bailey C, Bidigare R, Andersen R (2002) Phycol Res 50:31 46. Nakahara T, Yokochi T, Higashihara T, Tanaka S, Yaguchi T, Honda D (1996) J Am Oil Chem Soc 73:1421 47. Yokochi T, Honda D, Higashihara T, Nakahara T (1998) Appl Microbiol Biotechnol 49:72 48. Lewis T, Nichols P, McMeekin T (1999) Mar Biotechnol 1:580 49. Abril R, Garrett J, Zeller S, Sander W, Mast R (2003) Regul Toxicol Pharmacol 37:73 50. Hammond B, Mayhew D, Holson J, Nemec M, Mast R, Sander W (2001) Regul Toxicol Pharmacol 33:205 51. Hammond B, Mayhew D, Kier L, Mast R, Sander W (2002) Regul Toxicol Pharmacol 35:255

186

T. Matsunaga et al.

52. Hammond B, Mayhew D, Naylor M, Ruecker F, Mast R, Sander W (2001) Regul Toxicol Pharmacol 33:192 53. Hammond B, Mayhew D, Robinson K, Mast R, Sander W (2001) Regul Toxicol Pharmacol 33:356 54. Fan K, Chan F, Jones E, Vrijmoed L (2001) J Ind Microbiol Biotechnol 27:199 55. Sogawa K, Matsuda M, Okutani K (1998) J Mar Biotechnol 6:241 56. Sogawa K, Sumida T, Hamakawa H, Yamada T, Matsumoto K, Matsuda M, Oda H, Miyake H, Tashiro S, Okutani K (1998) Res Commun Mol Pathol Pharmacol 99:259 57. Sogawa K, Yamada T, Muramatsu Y, Sumida T, Hamakawa H, Oda H, Miyake H, Tashiro S, Matsuda M, Matsumoto K, Okutani K (1998) Res Commun Mol Pathol Pharmacol 99:267 58. Umemura K, Yanase K, Suzuki M, Okutani K, Yamori T, Andoh T (2003) Biochem Pharmacol 66:481 59. Yim J, Kim S, Ahn S, Lee H (2003) Biomol Eng 20:273 60. Burgess J, Iwamoto K, Miura Y, Takano H, Matsunaga T (1993) Appl Microbiol Biotechnol 39:456 61. Takeyama H, Iwamoto K, Hata S, Matsunaga T (1996) J Mar Biotechnol 3:244 62. Triemer R, Zimba P, Rowan M (2003) 55th Annual Meeting of the Society of Protozoologists 63. Takeyama H, Kanamaru A, Yoshino Y, Kakuta H, Kawamura Y, Matsunaga T (1997) Biotechnol Bioeng 53:185 64. Elhai J (1994) J Appl Phycol 6:177 65. Murphy R, Stevens S (1992) Appl Environ Microbiol 58:1650 66. Takeyama H, Takeda D, Yazawa K, Yamada A, Matsunaga T (1997) Microbiology 143(Pt 8):2725 67. Buzby J, Porter R, Stevens Jr S (1983) J Bacteriol 154:1446 68. Muro-Pastor A, Kuritz T, Flores E, Herrero A, Wolk CP (1994) J Bacteriol 176:1093 69. Matsunaga T, Takeyama H, Nakamura N (1990) Appl Biochem Biotechnol 24/25:151 70. Takeyama H, Nakayama H, Matsunaga T (2000) Appl Biochem Biotechnol 84–86:447 71. Kawaguchi R, Nagaoka T, Burgess J, Takeyama H, Matsunaga T (1994) Plasmid 72. Yang X, McFadden B (1993) J Bacteriol 175:3981 73. Akiyama H, Kanai S, Hirano M, Miyasaka H (1998) DNA Res 5:127 74. Elhai J, Wolk C (1988) Methods Enzymol 167:747 754 75. Vachhani A, Ramkumar K, Tuli R (1993) J Gen Microbiol 139:569 76. Chiang G, Schaefer M, Grossman A (1992) Physiol Biochem 30:315 77. Marraccini P, Bulteau S, Cassier-Chauvat C, Mermet-Bouvier P, Chauvat F (1993) Plant Mol Biol 23:905 78. Sode K, Tatara M, Takeyama H, Burgess J, Matsunaga T (1992) Appl Microbiol Biotechnol 37:369 79. Sode K, Tatara M, Ogawa S, Matsunaga T (1992) FEMS Microbiol Lett 99:73 80. Safferman R, Morris M (1963) Science 140:679 81. Sode K, Oozeki M, Asakawa K, Burgess JG, Matsunaga T (1994) J Mar Biotechnol 1:189 82. Bergh O, Borsheim K, Bratbak G, Heldel M (1989) Nature 340:467 83. Zhong Y, Chen F, Wilhelm S, Poorvin L, Hodson R (2002) Appl Environ Microbiol 68:1576 84. Shark K, Smith F, Harpending P, Rasmussen JL, Sanford J (1991) Appl Environ Microbiol 57:480 85. Matsunaga T, Sakaguchi T, Tadokoro F (1991) Appl Microbiol Biotechnol 35:651 86. Takeyama H, Kudo S, Sode K, Nakamura N, Matsunaga T (1991) Kobunshi Ronbunshu 48:319

Marine Microalgae 87. 88. 89. 90.

187

Apt KE, Kroth-Pancic P, Grossman A (1996) Mol Gen Genet 252:572 Dunahay T, Jarvis E, Roessler P (1995) J Phycol 31:1004 Falciatore A, Casotti R, Leblanc C, Abrescia C, Bowler C (1999) Mar Biotechnol 1:239 Geng D, Han Y, Wang Y, Wang P, Zhang L, Li W, Sun Y (2004) Acta Botanica Sinica 46:342 91. Yu R, Yamada A, Watanabe K, Yazawa K, Takeyama H, Matsunaga T, Kurane R (2000) Lipids 35:1061 92. Zaslavskaia L, Lippmeier J, Shih C, Ehrhardt D, Grossman A, Apt K (2001) Science 292:2073 93. Harker M, Hirschberg J (1997) FEBS Lett 404:129 94. Takahama K, Matsuoka M, Nagahama K, Ogawa T (2003) J Biosci Bioeng 95:302 95. Rocap G, Larimer F, Lamerdin J, Malfatti S, Chain P, Ahlgren N, Arellano A, Coleman M, Hauser L, Hess WR, Johnson Z, Land M, Lindell D, Post A, Regala W, Shah M, Shaw S, Steglich C, Sullivan M, Ting C, Tolonen A, Webb E, Zinser E, Chisholm S (2003) Nature 424:1042 96. Dufresne A, Salanoubat M, Partensky F, Artiguenave F, Axmann I, Barbe V, Duprat S, Galperin M, Koonin E, Le Gall F, Makarova K, Ostrowski M, Oztas S, Robert C, Rogozin IB, Scanlan D, Tandeau de Marsac N, Weissenbach J, Wincker P, Wolf YI, Hess W (2003) Proc Natl Acad Sci USA 100:10020 97. Palenik B, Brahamsha B, Larimer F, Land M, Hauser L, Chain P, Lamerdin J, Regala W, Allen E, McCarren J, Paulsen I, Dufresne A, Partensky F, Webb EA, Waterbury J (2003) Nature 424:1037 98. Scala S, Carels N, Falciatore A, Chiusano M, Bowler C (2002) Plant Physiol 129:993 99. Contreras-Flores C, Pena-Castro JM, Flores-Cotera LB, Canizares-Villanueva RO (2003) Interciencia 28:450 100. Lopezruiz JL, Garcia RG, Soledad M, Almeda F (1995) Aquat Eng 14:367 101. Benemann J (1992) J Appl Phycol 4:233 102. Jimmy RA, Kelly MS, Beaumont AR (2003) Aquaculture 220:261 103. Kaya V, de la Noue J, Picard G (1995) J Appl Phycol 7:85 104. Stirk WA, van Staden J (2001) S Afr J Bot 67:615 105. Matsunaga T, Takeyama H, Nakao T, Yamazawa A (1999) J Biotechnol 70:33 106. Borowitzka M (1995) J Appl Phycol 7:3 107. Lorenz RT, Cysewski GR (2000) Trends Biotechnol 18:160 108. Olaizola M (2003) Biomol Eng 20:459 109. Radmer RJ, Parker BC (1994) J Appl Phycol 6:93 110. (1994) The IEA Greenhouse Gas R&D Program, Report G/93/OE16B, Chemical Society of Japan 111. Torzillo G, Pushparaj B, Masojidek J, Vonshak A (2003) Biotechnol Bioprocess Eng 8:338 112. Richmond A (1992) J Appl Phycol 4:281 113. Chaumont D (1993) J Appl Phycol 5:593 114. Vonshak A, Boussiba S, Abeliovich A, Richmond A (1983) Biotechnol Bioeng 25:341 115. Duerr E, Molnar A, Sato V (1998) J Mar Biotechnol 6:65 116. Borowitzka M (1999) J Biotechnol 70:313 117. Borowitzka M (1996) J Mar Biotechnol 4:185 118. Janssen M, Tramper J, Mur LR, Wijffels RH (2003) Biotechnol Bioeng 81:193 119. Richmond A, Boussiba S, Vonshak A, Kopel R (1993) J Appl Phycol 5:327 120. Torzillo G, Carlozzi P, Pushparaj B, Montaini E, Materassi R (1993) Biotech Bioeng 42:891 121. Barbosa MJ, Hadiyanto, Wijffels RH (2004) Biotechnol Bioeng 85:78

188

T. Matsunaga et al.

122. Lee Y, Low C (1992) Biotechnol Bioeng 40:1119 123. Takano H, Takeyama H, Nakamura N, Sode K, Burgess J, Manabe E, Hirano M, Matsunaga T (1992) Appl Biochem Biotechnol 34/35:449 124. Barbosa M, Janssen M, Ham N, Tramper J, Wijffels R (2003) Biotechnol Bioeng 82:170 125. Barbosa M, Hoogakker J, Wijffels RH (2003) Biomol Eng 20:115 126. Kodama M, Ikemoto H, Miyachi S (1993) J Mar Biotechnol 1:21 127. Takano H, Takei R, Manabe E, Burgess JG, Hirano M, Matsunaga T (1995) Appl Microbiol Biotechnol 43:460 128. Takano H, Jeon J, Burgess J, Manabe E, Izumi Y, Okazaki M, Matsunaga T (1994) Appl Microbiol Biotechnol 40:946 129. Takano H, Matsunaga T (1995) Energ Convers Manage 36:697 130. Takano H, Manabe E, Hirano M, Okazaki M, Burgess J, Nakamura N, Matsunaga T (1993) Appl Biochem Biotechnol 39:239 131. Murakami M, Ikenouchi M (1997) Energ Convers Manag 38S:493

Adv Biochem Engin/Biotechnol (2005) 96: 189–218 DOI 10.1007/b135785 © Springer-Verlag Berlin Heidelberg 2005 Published online: 26 August 2005

Marine Enzymes Ghosh Debashish · Saha Malay · Sana Barindra · Mukherjee Joydeep (u) Environmental Science Programme and Department of Life Science & Biotechnology, Jadavpur University, 700 032 Kolkata, India [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract Marine enzyme biotechnology can offer novel biocatalysts with properties like high salt tolerance, hyperthermostability, barophilicity, cold adaptivity, and ease in largescale cultivation. This review deals with the research and development work done on the occurrence, molecular biology, and bioprocessing of marine enzymes during the last decade. Exotic locations have been accessed for the search of novel enzymes. Scientists have isolated proteases and carbohydrases from deep sea hydrothermal vents. Cold active metabolic enzymes from psychrophilic marine microorganisms have received considerable research attention. Marine symbiont microorganisms growing in association with animals and plants were shown to produce enzymes of commercial interest. Microorganisms isolated from sediment and seawater have been the most widely studied, proteases, carbohydrases, and peroxidases being noteworthy. Enzymes from marine animals and plants were primarily studied for their metabolic roles, though proteases and peroxidases have found industrial applications. Novel techniques in molecular biology applied to assess the diversity of chitinases, nitrate, nitrite, ammonia-metabolizing, and pollutant-degrading enzymes are discussed. Genes encoding chitinases, proteases, and carbohydrases from microbial and animal sources have been cloned and characterized. Research on the bioprocessing of marine-derived enzymes, however, has been scanty, fo-

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cusing mainly on the application of solid-state fermentation to the production of enzymes from microbial sources. Keywords Marine · Enzyme · Microorganisms · Molecular biology · Bioprocess

Abbreviations ArChi ATP BLAST bp c-AMP 3
,5
-CNP cbp cDNA chi cht DGGE DMS DMSP DNA GALT GDH h IPNV ISP kb kbp min m MABV MPa MUF-diNAG NAD(P) NADH nifH nir nit nos nrtP PAGE PAH PCR rRNA RuBisCO RdRp SBEs SSF β-HP

Arthrobacter chitinase Adenine triphosphate Basic Local Alignment Search Tool base pair 3
,5
-cyclic adenosine monophosphate cyclic nucleotide phosphodiesterase cellobiose phosphorylase complementary deoxyribonucleic acid chitinase genes N-acetyl-β-glucosaminidase denaturing gradient gel electrophoresis dimethyl sulfide dimethylsulfoniopropionate deoxyribonucleic acid galactose-1-phosphate uridylyltransferase glucose dehydrogenase hour infectious pancreatic necrosis virus iron-sulfur protein kilo base kilo base pair minute RNA messenger ribonucleic acid marine birnavirus mega pascal 4-methylumbelliferyl β-D-N, N
-diacetylchitobioside nicotinamide adenine dinucleotide (phosphate) hydrogenated nicotinamide adenine dinucleotide nitrogenase gene nitrite reductase gene Nitrosospira-like sequences nitrous oxide reductase gene nitrate/nitrite permease polyacrylamide gel electrophoresis polyaromatic hydrocarbon polymerase chain reaction ribosomal ribonucleic acid ribulose-1,5-bisphosphate carboxylase/oxygenase RNA-dependent RNA polymerase starch-branching enzymes solid-state fermentation β-hydroxypropionate

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1 Introduction The biological and chemical diversity of the marine environment has been the source of unique chemical compounds with the potential for industrial development as pharmaceuticals, cosmetics, nutritional supplements, molecular probes, enzymes, fine chemicals, and agrichemicals [1]. The oceans represent a virtually untapped resource for the discovery of even more novel compounds with useful activity. Although the commercial success stories in biotechnology are familiar, such stories in marine biotechnology are far less familiar and far fewer [2]. In the last decade there has been a continuous effort to learn more about the still largely unexplored realm of marine enzymes. In this review, the occurrence of enzymes from various marine sources will be described followed by application of molecular biology and finally the bioprocessing aspects of some marine biocatalysts. A marine enzyme may be a unique protein molecule not found in any terrestrial organism or it may be a known enzyme from a terrestrial source but with novel properties. Beside microorganisms like bacteria, fungi, and actinomycetes, many other marine organisms such as fishes, prawns, crabs, snakes, plants, and algae have also been studied to tap the arsenal of the marine world. Properties like high salt tolerance, hyperthermostability, barophilicity, cold adaptivity, and ease in large-scale cultivation are the key interests of scientists. These properties may not be expected in terrestrial sources as marine organisms thrive in habitats such as hydrothermal vents, oceanic caves, and some areas where high pressure and absence of light are obvious.

2 Source A survey of the literature of the past 10 years shows that the occurrence of marine enzymes has been most widely studied. Therefore, we begin the review with a description of various sources classified as marine microorganisms, marine animals, and plants. 2.1 Marine Microorganisms As marine microorganisms are very easy to tap, cultivate, identify (by molecular phylogenetic method), and bioprocess, they are of major interest to researchers worldwide. The symbiotic nature (microorganisms found associated with various marine sponges, corals, and other species) and their

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occurrence in extreme environments (extremophiles) like hydrothermal vents have also been areas of recent research and are therefore reviewed in this article. 2.1.1 Extremophiles Diversa, an American company engaged in the application of microbial biodiversity in the biotechnology industry, recognized that thermophilic enzymes from vents with temperatures of 350–400 ◦ C are stable protein molecules. Even if they are to be used at mild temperatures, they remain active far longer than regular enzymes. They also resist the destabilizing effects of organic chemicals used in industrial downstream processes. A recent US National Academy of Sciences report noted that in 1993, world enzyme sales equaled US$ 1 billion, a market that was expected to grow about 10% each year. Enzymes from extremophiles will constitute a part of this market [3]. A second generation of thermostable polymerase chain reaction (PCR) enzymes has already been harvested from bacteria living near thermal vents on the ocean floor and is marketed as Vent and Deep Vent polymerases [4]. Similarly, psychrophilic enzymes can be useful for commercial laundry detergents as consumers can wash clothes in cold rather than hot water, which could significantly reduce power consumption. Although such applications have been projected, current literature, however, is limited to the occurrence of metabolic enzymes in psychrophilic organisms [5, 6]. Major recent advances in cold deep sea biotechnology have come in the form of continuing discoveries of novel microorganisms, with unexpected genetic diversity and new natural products including enzymes of potential relevance to human health or environmental bioremediation [7, 8]. Continuing explorations of submarine hydrothermal vent environments have yielded new hyperthermophiles and more evidence of pressure-regulated operons and elevated hydrostatic pressure stabilization of cells and enzymes at high temperatures. This section is broadly divided into three parts, describing enzymes from thermophiles, psychrophiles, and piezophiles. Michels et al. [9] reported the properties of a hyperthermophilic, barophilic protease from Methanococcus jannaschii, the first protease to be isolated from an organism adapted to a high-pressure-high-temperature environment. A protease was isolated and purified from the supernatant of a culture of hyperthermophilic archaebacteria Pyrococcus abyssi by Dib et al. [10]. A novel intracellular serine proteinase (pernilase) was identified from a marine aerobic hyperthermophilic archaeon Aeropyrum pernix having enzyme half-lives of 85 min at 100 ◦ C and 12 min at 110 ◦ C by Chavez et al. [11]. After extensive investigation of shallow water and deep sea hydrothermal vents for the isolation of hyperthermophilic microorganisms, mostly Pyrococcus sp. and Thermococcus sp., thermostable hydrolytic enzymes were character-

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ized for potential applications. Optimal growth conditions were analyzed to determine final yields and metabolic rates; the effects of temperature and hydrostatic pressure on cell replication and enzymatic activities were also investigated [12]. Microorganisms growing above 60 ◦ C isolated from deep sea hydrothermal vents were screened for amylolytic activity. Nine archaea and one thermophilic bacterium were selected for the determination of thermostability and pH optima. Pullulanase, glucosidase, and amylase activities were detected in four archaeal strains related to the genus Thermococcus [13]. Brown and Kelly [14] purified extracellular pullulanases from cell-free culture supernatants of the marine thermophilic archaea Thermococcus litoralis and Pyrococcus furiosus. The enzymes from T. litoralis and P. furiosus appeared to represent highly thermostable amylopullulanases, versions of which, however, have been isolated from less thermophilic organisms. Gantelet and Duchiron isolated the extremely thermophilic archaeon T. hydrothermalis from a deep sea hydrothermal vent in the East Pacific Rise, which produced an extracellular pullulanase [15]. The chitinoclastic enzyme system of T. chitonophagus, isolated from a hydrothermal vent site off the Mexican west coast, was oxygen-stable, cell-associated, and inducible by chitin [16]. The membrane-bound hydrogenase from a marine hydrogen-oxidizing bacterium Hydrogenovibrio marinus was characterized by Nishihara et al. as highly oxygen tolerant, extremely thermophilic, and thermostable in its membrane-bound form [17]. The pyruvate carboxylase of Methanococcus jannaschii was purified and expressed in cells grown without an external source of biotin [18]. Four different tungsten-containing enzymes have been purified from P. furiosus that oxidize aldehydes of various types and are thought to play primary roles in the catabolism of sugars or amino acids [19]. Turning to psychrophiles and psychrotolerant organisms, recent studies have elucidated that although microorganisms produce various psychrophilic enzymes in order to carry out metabolism efficiently under cold conditions, enzymes studied so far are heat stable. A psychrophile from Antarctic seawater identified as Cytophaga sp., which grows optimally at 15 ◦ C but cannot grow above 30 ◦ C, produces a variety of NAD(P)-dependent dehydrogenases, among which alcohol dehydrogenase and aldehyde dehydrogenase are thermostable, that has been reported by Soda et al. [20]. Alanine dehydrogenase, malate dehydrogenase, and glutamate dehydrogenase were detected in extracts from a psychrophilic bacterium, closest to Shewanella isolated from a sea urchin off the Icelandic coast by Irwin et al. [21]. An extracellular serine peptidase was purified from the culture supernatant of a sub-Arctic psychrophilic bacterium. It was a remarkably stable enzyme from a psychrophilic microorganism, remaining active after 1 week at 20 ◦ C and after five freeze-thaw cycles [22]. Kazuoka et al. [23] reported that Cytophaga sp. produces aspartase abundantly. The enzyme showed higher pH and thermal stabilities than that from mesophiles. Two enzymes from Colwellia maris—

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isocitrate lyase having maximum activity at 20 ◦ C but rapidly inactivated at temperatures above 30 ◦ C and malate synthase having optimum temperature of activity of 45 ◦ C—were discovered [24]. Possible explanations for the presence of a cold-active enzyme in Fibrobacter succinogenes, a mesophile, are that cold-active enzymes are more broadly distributed and that lateral transfer of the gene from a psychrophile occurred or that F. succinogenes originated from the marine environment [25]. Marine psychrophilic bacteria with β-1,3-glucanase activities have been isolated, and the strain with the highest glucanase activity has been taxonomically identified and the enzyme was purified [26]. A heat-labile β-lactamase similar to highly specialized cephalosporinases from pathogenic mesophilic bacteria has been purified by Feller et al. [27] from the cold-adapted psychrophile Psychrobacter immobilis. A few studies have been reported on enzymes from piezophilic microorganisms. Konisky et al. [28] noted that the application of 50 MPa pressure did not increase the thermostabilities of adenylate kinases purified from four related mesophilic and thermophilic marine methanogens in contrast to the evidence on hyperbaric stabilization of enzymes from deep sea thermophiles. Researchers investigated the respiratory chain system of a deep sea barophilic bacterium Shewanella sp., and a novel heme c containing quinol oxidase was purified from cells grown at 60 MPa pressure. A study of its properties by Qureshi et al. suggested the presence of two kinds of respiratory chains regulated in response to pressure in this bacterium [29]. 2.1.2 Marine Symbiotic Microorganisms Marine invertebrates and their cultivatable bacterial symbionts have become a focal point of marine natural product research. Basic research on some of these associations have resulted in the biotechnological development of these valuable resources. A bacterial protease was isolated from a marine shipworm and tested in cleansing formulations [30]. Bacterial mats that grow on whales have been found to be a rich source of lipases and esterases, two classes of enzymes important to the industry [5]. Marine microorganisms often survive as intracellular or extracellular symbionts, and their hosts are mostly marine animals (vertebrates or invertebrates), which are first described in this section, followed by one example of a plant host. The deep sea tube worm Riftia pachyptila (Vestimentifera) from hydrothermal vents lives in an intimate symbiosis with a sulfur-oxidizing bacterium, and investigations indicate that the animal is fully dependent on the symbiont for the de novo biosynthesis of pyrimidines [31]. The arginine biosynthetic enzymes are present in all the tissues of the worm and in the bacteria [32]. Scientists present microscopical and enzymatic (presence of methanol dehydrogenase) evidence that methylotrophic bacteria occur as intracellular symbionts in a new species of mytilid mussel discovered at the

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mid-Atlantic ridge hydrothermal vents [33]. Pseudoalteromonas sp. isolated from the alimentary tract of Antarctic krill Thyssanoessa macrura synthesizes an intracellular cold-adapted β-galactosidase [34]. Mohapatra [35] reported the isolation of carboxymethylcellulase from a Bacillus sp. associated with the marine sponge Axinella sp. The marine sponge Spirastrella sp. was found to be an interesting host organism as it harbored a Mucor sp. producing a novel amylase [36], a Micrococcus sp. producing urethenase [37], and an acetylcholinesterase producing bacterium Arthrobactor ilicis [38]. An extracellular protease from the marine bacterium Sphingomonas paucimobilis isolated from the stomach of Antarctic krill Euphausia superba Dana was purified and characterized [39]. Vibrio fischeri, a marine bacterium that forms a bioluminescent symbiosis with certain fish and squids, exhibits the unusual attribute of growth on 3
,5
-cyclic adenosine monophosphate (c-AMP), apparently through the activity of a 3
,5
-cyclic nucleotide phosphodiesterase (3
,5
-CNP) with exceptionally high activity [40], which, due to its unusual location in the periplasm, allows this symbiotic bacterium to utilize extracellular 3
,5
-cyclic nucleotides (e.g., c-AMP) as sole sources of carbon, nitrogen, and phosphorus [41]. A Streptomycetes sp. isolated from the prawn Penaeus indicus showed good L-asparaginase activity [42]. A Bacillus sp. [43] and a Mucor sp. [44] associated with the intertidal marine alga Sargassum sp. also were found to be L-asparaginase producers. The only example of a plant host is Laminaria fronds that house a bacterium Alteromonas sp. producing extra- and intracellular alginate lyases and utilizes alginate as its sole carbon source [45]. 2.1.3 Microorganisms from Marine Sediment and Seawater Near shore sediments, deep sea sediments, and seawater have been easily reachable to marine biotechnologists, and therefore reports of marine enzymes from these sources have been numerous over the past decade. Among them, proteases, carbohydrases, and peroxidases have been the most cited ones. Some of them have found commercial applications. The extracellular proteases are of particular importance and can be used in detergents and industrial cleaning applications, such as in cleaning reverse osmosis membranes. Vibrio spp. have been found to produce a variety of extracellular proteases. V. alginolyticus produces six proteases, including an unusual detergent-resistant, alkaline serine exoprotease. This marine bacterium also produces collagenase, an enzyme with a variety of industrial and commercial applications [46]. BAL 31 Nuclease, manufactured by the Japanese firm TaKaRa, is produced by the marine bacterium Alteromonas espejiana BAL 31, an endonuclease specific to single-stranded deoxyribonucleic acid (DNA) (activity I) and also has exonuclease activity (activity II).

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Hydrolases Shibata et al. discovered a novel metalloproteinase, almelysin, which has high activity at low temperatures, and another proteinase from the culture supernatant of a marine bacterium, Alteromonas sp. [47]. A protamine-degrading marine bacterium was isolated from marine soil and identified as Aeromonas salmonicida subsp. [48]. Alteromonas sp. secretes a metalloprotease involved in the chitin degradation system of the strain [49]. Rath and Herndl [50] reported that marine snow of the northern Adriatic Sea showed β-D-glucosidase activity. Results obtained by Arrieta and Herndl [51] with natural bacterial communities analyzed by capillary electrophoresis zymography from the coastal North Sea suggest that the diversity of β-glucosidases in the marine environment might be much higher than previously observed. In order to identify strains with α-1,4- and 1,6-glucosidase enzymes with potential uses in shrimp feed production, Bacillus subtilis was isolated from marine environments by Arellano–Carbajal and Olmos– Soto [52]. Optimal conditions for growth of a marine fungus Chaetomium indicum and for biosynthesis of β-1,3-glucanase were determined by Burtseva et al. [53]. The extracellular enzymatic activity of a mixed culture of anaerobic marine bacteria enriched on pullulan was studied by Arnosti and Repeta [54]. A β-mannanase from Vibrio sp. was purified by Tamaru et al. [55]. A marine bacterial strain isolated from the Bay of San Vicente, Chile, identified as Alteromonas sp., produced high levels of an extracellular agarase in the presence of agar [56]. A marine bacterial strain of genus Vibrio that decomposes the cell walls of some seaweed, including a Laminaria sp. and Undaria pinnatifida, was isolated from seawater by Sugano et al. [57]. A novel enzyme, α-neoagarooligosaccharide hydrolase, which hydrolyzes the α-1,3 linkage of neoagarooligosaccharides to yield agaropentaose and D-galactose, was isolated from the marine bacterium Vibrio sp. and characterized by Sugano et al. [58]. The phenotypic and agarolytic features of a marine bacterium Pseudoalteromonas antarctica that was isolated from the southern Pacific coast was investigated. It produced a diffusible agarase that caused agar softening around the colonies [59]. β-agarase, from marine bacterium Bacillus cereus, was purified and characterized and gave a single band on polyacrylamide gel electrophoresis (PAGE) with activity staining [60]. Another marine bacterium degraded numerous complex carbohydrates, such as agar, chitin, and alginate, with an agarase system that consisted of at least three enzymes, β-agarase I, β-agarase II, and α-agarase, which acted in concert to degrade polymeric agar to D-galactose and 3,6-anhydro-L-galactose [61]. The degradation of chitin, the most abundant polymer in marine environment, was studied. Chitinase from the marine bacterium Alteromonas sp. was purified [62]. V. harveyi was found to have a higher growth rate and more chitinase activity when grown on β-chitin (isolated from squid pen) than on α-chitin (isolated from snow crab) probably because of the more open

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structure of β-chitin. When exposed to different types of chitin, V. harveyi excreted several chitin-degrading proteins into the culture media [63]. Marine bacterial strains of Cytophaga diffluens, C. hutchinsonii, Pseudomonas sp. (fluorescent) and V. fluvialis were examined for cellulase production using four media by Vaidya et al. [64], and Araki et al. [65] reported that β-1,3xylanase was purified to gel electrophoretic homogeneity from a cell-free culture fluid of Vibrio sp. Phosphate solubilizing bacteria belonging to Bacillus sp. were found to be highly adaptive and therefore can significantly contribute to the phosphate economy of the marine environment [66]. A marine Vibrio sp. producing a particularly heat-labile alkaline phosphatase was isolated by Hauksson et al. from North Atlantic coastal waters [67]. Constitutive amidase with broad substrate specificity from a number of deep sea Actinomycetes were reported by Brandão and Bull [68]. Oxidoreductases Chloroperoxidase isolated from the marine fungus Caldariomyces fumago is unique among the peroxidases because it contains a cysteinic thiolate as the fifth axial ligand of the heme instead of the imidazole ligand. This enzyme is unusually versatile: it catalyzes not only the reactions typical of peroxidases but also those of catalases and monooxygenases, and it is also almost unique in catalyzing halogenation reactions (except fluorination) in the presence of halide ions and H2 O2 [69]. A basidiomycetes fungus Flavodon flavus isolated from decaying sea grass from a coral lagoon off the west coast of India produced three major classes of extracellular lignin-modifying enzymes: manganese-dependent peroxidase, lignin peroxidase, and laccase [70]. A yellow pigmented marine bacterium was isolated by Francis et al. from surface sediments of San Diego Bay based on its ability to oxidize soluble Mn (II) to insoluble Mn (III, IV) oxides [71]. Manganese-dependent peroxidase activities detected in culture supernatants of Debaryomyces polymorphus, Candida tropicalis, and Umbelopsis isabellina were responsible for color removal of synthetic dyes by enzymatic biodegradation [72]. Three constitutive forms of superoxide dismutase (iron, copper/zinc, and an unidentified form) activity have been demonstrated in the marine cyanobacterium Synechococcus sp. by Chadd et al. [73]. The cytosolic form of Cu-Zn superoxide dismutase has been isolated from the marine yeast Debaryomyces hansenii [74]. A novel glucose dehydrogenase (GDH) from a marine bacterium Cytophaga marinoflava was isolated by Tsugawa et al. [75] from its membrane fraction. The GDH can react under high salinity. Another novel GDH that reacts with a clinical marker of diabetes was purified by the same group [76] from a soluble fraction of a marine Gram-negative bacterium identified as a Deleya sp. Nitrite reductase was purified to electrophoretic homogeneity from the soluble extract of the marine denitrifying bacterium Pseudomonas nautica [77].

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Lyases Dimethyl sulfide (DMS), the most abundant volatile sulfur compound emitted from oceans, is formed primarily by the action of dimethylsulfoniopropionate (DMSP) lyase, which cleaves DMSP, an algal osmolyte, to equimolar amounts of DMS and acrylate. D’Souza and Yoch [78] reported the isolation and purification of DMSP lyase. The soluble enzyme was purified to electrophoretic homogeneity from a facultatively anaerobic Gram-negative rod-shaped marine bacterium identified as an Alcaligenes sp., a salt marsh bacterial isolate. The production of DMSP lyase from this organism and a marine strain, Pseudomonas doudoroffii, were induced at optimum rates by DMSP and vigorous aeration [79]. DMSP lyase was also isolated from the sulfate-reducing bacterium by Jansen and Hansen [80]. It was demonstrated in an α-subclass of Proteobacteria in marine bacterioplankton community that DMSP was taken up and metabolized by an intracellular DMSP lyase and acrylase. Added acrylate was β-hydroxylated on (or near) the cell surface to β-hydroxypropionate (β-HP), which accumulated briefly and was then taken up by cells. DMSP, acrylate, and β-HP all induced DMSP lyase activity [81]. Fucobacter marina, a marine bacterial strain isolated by Sakai et al., produced extracellular sulfated fucoglucuronomannan lyase [82]. Highly active constitutive nitrile hydratase with broad substrate specificity from several deep sea Actinomycetes were reported by Brandão and Bull (2003) [68]. Ligases Glutamine synthetase in cells of the marine diazotrophic cyanobacterium Trichodesmium thiebautii was investigated by Carpenter et al. [83]. The physiological regulation of glutamine synthetase in the axenic cyanobacterium Prochlorococcus sp. was studied, and the unusual responses to darkness and nitrogen starvation could reflect adaptation mechanisms of Prochlorococcus sp. for coping with a light- and nutrient-limited environment [84]. Other enzymes Recent reports of other marine microbial enzymes belonging to the above categories obtained from sediment and seawater include lycopene k-cyclase [85], L-serine dehydratase [86], polyhydroxybutyrate depolymerase [87], quinol oxidase [88], and arylsulphatase [89]. Our research group has, for the first time, conducted surveys in the deltaic Sundarbans, the world’s largest tidal mangrove forest located in the delta of the Ganges, Brahmaputra, and Meghna rivers on the Bay of Bengal. We have isolated several sediment-dwelling marine bacteria producing salt-tolerant and thermostable protease, esterase, nitrilase, ribonuclease, L-asparaginase, and L-glutaminase [90].

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2.2 Marine Animals In the past decade enzymes from marine animals have been explored for their metabolic roles and potential industrial applications. Some products are already available on the market and some are going through clinical trials. The Sunlife Corporation (USA) marketed Penzim; the active ingredient penzyme is a digestive protease trypsin from the North Atlantic cod. It is a very powerful psychrophilic proteolytic enzyme. Neptune Technologies & Bioresources introduced Neptune Krill Enzymes, which has natural powerful digestive enzymes like proteases, phosphatases, and phosphohydrolases combined with peptides. Another product, Neptune Aquatein, is the dry fraction remaining after the extraction of Neptune Krill Oil. Research has shown that Aquatein enzymes are lipases, phospholipases, alkaline phosphatase, acid phosphatase, esterase, trypsin, phosphohydrolase, glucoronidase, glucosidase, proteases, hyalurinases, and nucleases. Participation of academia and industry in marine enzyme biotechnology has been exemplified in the development of a cold active lysozyme-chlamysin with antimicrobial activity. Nilsen et al. have isolated this enzyme from the viscera of the marine bivalve Chlamys islandica [91]. Fiskeriforskning, a Norwegian biotechnology firm, has also isolated this enzyme [92], and the encoding complementary DNA (cDNA) gene that actuates the enzyme production in scallops has been analyzed by Nilsen and Myrnes [93]. Table 1 presents an overview of biotechnological applications and metabolic functions of enzymes isolated from marine animals.

Table 1 Functions and applications of enzymes isolated from various marine animal sources Enzyme Class

Enzyme

Source

Biotechnological applications/ Reference metabolic functions

Marine sponge Spheciospongia vesparia

Degrade casein, hide powder azure, synthetic substrates

[94]

Cathepsins

Marine sponge Geodia cydonium

Major digestive protease in sponges

[95]

Protease

Marine crab, Scylla serrata

Collagenolytic metalloprotease closely resembles metalloproteases of vertebrates

Sparus aurata, Scophthalmus maximus and Sebastes mentella

Digestive enzymes in marine fishes

Hydrolase Protease

Amylase

[96]

[97]

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Table 1 (continued) Enzyme class

Transferase

Oxidoreductase

Enzyme

Source

Biotechnological applications/ Reference metabolic functions

Cholinesterases

Bivalve Mytilus Biomarker for aquatic edulis, Mytilus pollution galloprovincialis, Corbicula fluminea

[98]

AMPdeaminase

Teleost sea scorpion, Scorpaena porcus

Purine nucleotide metabolism

[99]

Na, K-ATPase

Spiny lobster Palinurus elephas

Generation of osmolyte gradient

[100]

Hyaluronidase

Venom of the Only marine hyaluronidase stonesh, Synanceja horrida

[101]

ATP N-glycosidase

Marine sponge Conversion of adenosineAxinella polypoides 5-triphosphate into adenine and ribose-5-triphosphate

[102]

α-Nacetylgalactosaminidase

Sea squirt

Structural analyses of the carbohydrate epitopes

[103]

Citrate synthase, Pyruvate kinase

Northern krill, Meganyctiphanes norvegica

Metabolic key enzymes, adaptive properties under different thermal conditions

[104]

TransRed sea bream glutaminase liver

Transfer of amine groups

[105]

Glutathione Sea bass (DicenS transferase trarchus labrax) liver cytosol

Novel glutathione S transferases belonging to θ and α classes

[106]

cAMPdependent protein kinase

Marine periwinkle, In reversible protein [107] Littorina littorea phosphorylation, aerobic-anaerobic transitions

Dehalogenating peroxidase

Terebellid polychaete, Amphitrite ornata

Removal of anthropogenic or biogenic haloaromatic compounds

[108]

Catalase, Superoxide dismutase

Marine mussel, Mytilus edulis L.

Antioxidant enzyme, Potential use in toxicological studies

[109]

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Table 1 (continued) Enzyme class

Enzyme

Source

Biotechnological applications/ Reference metabolic functions

Monooxygenase

Sea bass (Dicentrarchus labrax)

Biomarkers of polycyclic aromatic hydrocarbon exposure

[110]

Phenoloxidase

Marine mussel Perna viridis

Oxidation of phenolic substrates

[111]

20 βJapanese eel hydroxysteroid dehydrogenase

Key steroidogenic enzyme

[112]

Lyase

Phospholipases A2

Novel purification method

[113]

Other

Glucose Marine borer metabolism Bankia setacea enzymes

Metabolic enzyme

[114]

Sea snake Hydrophis cyanocinctus venom

2.3 Marine Plants Marine plants, especially marine algae, in recent years have been appealing candidates for bioprospecting of novel enzymes. Many commercially important enzymes have been isolated from marine phytoplanktons. Research has demonstrated the presence of unique haloperoxidases (e.g., vanadium bromoperoxidase with a high degree of stability to thermal and organic solvent denaturation) in algae. These enzymes could become valuable products because halogenation is an important process in the chemical industry. Japanese researchers have developed methods to induce a marine alga to produce large amounts of the enzyme superoxide dismutase, which is used in enormous quantities for a range of medical, cosmetic, and food applications. Table 2 shows the recent trends in research on the biotechnology of enzymes isolated from marine plants.

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Table 2 Functions and applications of enzymes isolated from various marine plant sources Enzyme class

Enzyme

Source

Biotechnological applications/ Reference metabolic functions

Oxidoreductase

Iodoperoxidase

Marine diatom cultures

Iodine incorporating enzyme [115]

Vanadiumbromoperoxidase

Lithophyllum yessoense

Potential substitute for catalase

Bromoperoxidase

Marine algae Regiospecific bromoperAscophyllum nod- oxidative oxidation osum and Corallina of 1,3-di-tert-butylindole officinalis

[117]

Nitrate reductase

Eelgrass, Zostera marina L.

[118]

Hydrogenase Marine green alga, New source Chlorococcum littorale

[119]

Luciferase

Marine dinoflagel- Enzymatic oxidation lates, Lingulodinium of luciferin polyedrum and Pyrocystis lunula

[120]

Xanthine oxidoreductases

Gonyaulax polyedra Role in circadian control

[121]

Marine green alga, fibrinolytic enzyme Codium divaricatum

[122]

Cholinesterase

Gracilaria corticata (Rhodophyta)

Metabolic enzyme

[123]

ATPase

Zostera marina

Salt-tolerant metabolic enzyme

[124]

Urease

Aureococcus anophagefferens, Prorocentrum minimum and Thalassiosira weissflogii

Conversion of urea to ammonia

[125]

Carbonic anhydrase

Marine diatom Phaeodactylum tricornutum

Hydration of CO2 and the dehydration of bicarbonate

[126]

Hydrolase Protease

Lyase

Key enzyme in nitrate assimilation Metabolic enzyme

[116]

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Table 2 (continued) Enzyme class

Ligase

Enzyme

Source

Biotechnological applications/ Reference metabolic functions

Myrcene synthase

Marine red alga Ochtodes secundiramea

Produces myrcene from geranyl diphosphate

Glutamine synthetase

Emiliani huxleye

Activity related with [128] light and nitrogen availability

[127]

Transferase Ribulose-1,5- Marine bisphosphate phytoplankton carboxylase/ oxygenase

Calvin cycle enzyme, Metabolic enzyme

[129]

Isomerase Polyenoic fatty acid isomerase

Conversion of arachidonic acid

[130]

Marine alga Ptilota filicina J

3 Molecular Biology A survey of the recent literature shows that molecular biology tools have been applied for assessing the biodiversity of marine enzymes, cloning, and characterizing enzyme genes. Investigations in this sphere of research should have immense commercial applications. The cDNA encoding silicatein, the first silica-synthesizing enzyme from the sponge Suberites domuncula, was used as a probe to study the potential role of silicate on the expression of the silicatein gene. It was found that after increasing the concentration of soluble silicate in the seawater medium, this gene is strongly upregulated [131]. This discovery has led to the European Community-funded project to develop new routes for the structure-controlled biofabrication of silica nanostructure materials for biosensors, biomedical uses, and biosemiconductors by diatoms and siliceous sponges and for the industrial and medical application of the enzymes involved in synthesis and dissolution of biogenic silica. The gene encoding Pyrolase 160 (Diversa), a broad-spectrum β-mannanase (from the deep sea hydrothermal vent), was discovered via expression screening and then inserted into a microorganism known to be an efficient and safe expression host.

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3.1 Molecular Methods to Assess Diversity Cloning ribosomal ribonucleic acid (rRNA) genes from mixed microbial assemblages is done to determine the phylogenetic identity of population constituents. Such cultivation-independent molecular phylogenetic surveys have revealed an astounding number of novel phylogenetic lineages [132]. Recent advances include the recovery of greater overall amounts of DNA in environmental DNA libraries. Rapid progress in high-throughput screening, sequencing, and robotics have also greatly facilitated a more thorough analysis of the recovered clones. These technological advances are vastly improving the economic and technical feasibility of cloning, screening, and sequencing large numbers of clones derived from natural environments. There has been a good deal of interest in recovering microbial DNA from soil, with most studies concentrating on bioprospecting for drugs, enzymes, and other natural products. This type of approach has been in use now for nearly a decade in the biotechnology industry [133]. Nowadays genetic engineers not only catalog rRNAs (or other single genetic loci), but also determine large portions of the genomic content found within naturally occurring microbial communities. Different opportunities have become perceptible through this new approach. Bioprospecting, characterization of uncultivated microbes, and microbial population genomics are advancing by its application [134]. In the early 1990s, molecular biology was fortified with the use of thermostable DNA polymerases and the PCR [135], which became a major tool for phylogenetic diversity study of single genetic loci, especially rRNA genes [136]. 3.1.1 Oxidoreductases Genetic heterogeneity of denitrifying bacteria in sediment samples from Puget Sound and two sites on the Washington continental margin was studied by PCR approaches by amplifying nitrite reductase genes (nirK and nirS). The findings demostrated a very high diversity of nir sequences within small samples and that these novel nir clusters, some very divergent from known sequences, were not known in cultivated denitrifiers [137]. Grüntzig et al. [138] used real-time PCR to quantify the denitrifying nitrite reductase gene of Pseudomonas stutzeri, a functional gene of biogeochemical significance from Puget Sound and from the Washington ocean margin. These results suggested that P. stutzeri may not be a dominant marine denitrifier. Diversity of the nitrous oxide reductase gene (nosZ) was examined by Scala and Kerkhof in sediments obtained from the Atlantic Ocean and Pacific Ocean continental shelves. Phylogenetic analysis illustrated three major clusters of nosZ genes with little overlap between environmental and culture-based groups. The two non-culture-based gene clusters generally cor-

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responded to the sampling location, implying that denitrifier communities may be restricted geographically [139]. Nitrogenase gene (nifH) sequences amplified directly from oceanic waters showed that the open ocean contains more diverse diazotrophic microbial populations and more diverse habitats for nitrogen fixers than previously observed by classical culture-based techniques. Nitrogenase genes derived from unicellular and filamentous cyanobacteria, as well as from the α and γ subdivisions of the class Proteobacteria, were found in both the Atlantic and Pacific oceans [140]. With the aim of testing the hypothesis that biological nitrogen fixation plays an important role in nitrogen cycling in the subseafloor associated with unsedimented hydrothermal vents, degenerate PCR primers were designed to amplify the nitrogenase protein gene nifH from hydrothermal vent fluid by Mehta et al. Potential nitrogen fixers were encountered in anaerobic Clostridia, and sulfate reducers included Proteobacteria and divergent Archaea. All of the nifH genes from the deep seawater sample were most closely related to the thermophilic, anaerobic archaeon Methanococcus thermolithotrophicus denoting that at least two sources contribute to the diverse assemblage of nifH genes detected in hydrothermal vent fluid, first, nifH genes from an anaerobic hot subseafloor and second, nifH genes from cold oxygenated deep seawater [141]. The spatial distribution and diversity of ammonia-oxidizing bacteria of the β subdivision of the class Proteobacteria (ammonia oxidizers) in the Arctic Ocean and Western Arctic Ocean were determined by Bano and Hollibaugh. The presence of ammonia oxidizers was detected by PCR amplification of 16S rRNA genes using a primer set specific for this group of organisms. Analysis of nitA–nitB (Nitrosospira-like sequences) PCR product by nested PCR denaturing gradient gel electrophoresis (DGGE) showed the presence of a dominant, ubiquitous ammonia oxidizer in the Arctic Ocean basin. 22% of the samples contained additional major bands. These samples were restricted to the areas influenced by Pacific inflow. The nucleotide sequence of the 1.1-kb nitA–nitBPCR product grouped with sequences designated “Group 1-marine Nitrosospira-like sequences”. Results connote that the Arctic Ocean β-proteobacterial ammonia oxidizers have low diversity and are dominated by marine Nitrosospira-like organisms. Diversity appeared to be higher in the Western Arctic Ocean [142]. The diversity of ammoniaoxidizing bacteria in aquatic sediments of the Pacific Northwest was studied by retrieving ammonia monooxygenase and methane monooxygenase gene sequences by Nold et al. Methanotrophs dominated freshwater sediments, while β-proteobacterial ammonia oxidizers dominated marine sediments. These results show that γ -Proteobacteria such as Nitrosococcus oceani are minor members of marine sediment ammonia-oxidizing communities [143]. The diversity of a gene encoding a key ring cleaving enzyme of the β-ketoadipate pathway, dioxygenase, amplified from bacterial communities associated with decaying Spartina alterniflora as well as from enrichment

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cultures with aromatic substrates (p-hydroxybenzoate, anthranilate, vanillate, and dehydroabietate) was investigated by Buchan et al. 52% of the clones could be assigned to the Roseobacter group of the class α-Proteobacteria abundant in coastal ecosystems. Another 6% of the clones matched genes retrieved from isolates belonging to the genera Acinetobacter, Bacillus, and Stappia, and 42% of the clones could not be assigned to any cultured bacterium based on sequence identity [144]. Degenerate primers and the PCR were used to isolate a portion of a naphthalene dioxygenase iron-sulfur protein (ISP) gene from a new bacterium, Neptunomonas naphthovorans. A phylogenetic analysis of polycyclic aromatic hydrocarbon dioxygenase ISP-deduced amino acid sequences showed that the genes isolated were distantly related to the genes encoding naphthalene dioxygenases of Pseudomonas and Burkholderia. 16S rDNA-based phylogenetic analysis placed these bacteria in the γ -3 subgroup of the Proteobacteria, most closely related to members of the genus Oceanospirillum. However, morphologic, physiologic, and genotypic differences between the new isolates and the Oceanospirilla justified the creation of a novel genus and species, N. naphthovorans [145]. Two distinct Cycloclasticus partial polyaromatic hydrocarbon (PAH) dioxygenase ISP gene sequences were PCR amplified from samples collected from Puget Sound and the Gulf of Mexico by Geiselbrecht et al. Cycloclasticus species appeared to be numerically important and widespread PAH-degrading bacteria in both Puget Sound and the Gulf of Mexico [146]. 3.1.2 Hydrolases PCR primers were designed based on chitinase (chi) genes in four γ -Proteobacteria in the families Alteromonadaceae and Enterobacteriaceae (Group I chitinases) and used to explore the occurrence and diversity of these chi genes in cultured and nonculturable marine bacteria from coastal Pacific Ocean and estuarine Delaware Bay bacterioplankton. The PCR results from 104 bacterial strains indicated that this type of chi gene occurs in two major groups of marine α and γ Proteobacteria, but not the Cytophaga-Flavobacter group [147]. To examine the ecology and evolution of microbial chitinases, especially the chitin-binding domain, one of the chi genes (chiA) from the marine bacterium Vibrio harveyi was analyzed by Vsitil and Kirchman [148]. Cottrell et al. identified representative and abundant chi genes from uncultivated marine bacteria and constructed libraries of genomic DNA isolated from coastal and estuarine waters. The libraries were screened for genes encoding proteins that hydrolyze a fluorogenic analog of chitin, 4-methylumbelliferyl β-D-N,N
-diacetylchitobioside (MUF-diNAG). The number of clones detected with the plaque assay was consistent with estimates of the portion of culturable bacteria that degrade chitin. Results signified that

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culture-dependent methods do not greatly underestimate the portion of marine bacterial communities capable of chitin degradation [149]. Urea appears to be a major nitrogen resource in the sea, but little molecular information exists about its utilization by marine organisms. Oligonucleotide primers were used to amplify a conserved fragment of the urease coding region from marine cyanobacteria. A 5.7-kbp region of the genome of the unicellular marine cyanobacterium Synechococcus sp. was cloned, and genes encoding three urease structural subunits and four urease accessory proteins were sequenced and identified by homology. The urease had a predicted subunit composition typical of bacterial ureases, but the organization of the urease genes was unique. Biochemical characteristics of the urease enzyme were consistent with the predictions of the sequence data. Physiological data and sequence analysis both suggested that the urease operon may be nitrogen regulated [150]. The cDNA encoding the putative prolidase was cloned from a library of demosponge Suberites domuncula by Wiens et al. Two different forms of cDNAs were identified, coding for the putative polypeptides of molecular mass 55 805 Da and 51 684 Da. Phylogenetic analysis revealed that the sponge prolidase branches off first from the common ancestor of metazoan prolidases and later from the yeast prolidase; only distantly related are the bacterial enzymes [151]. A set of marine psychrophilic bacteria has been surveyed in a PCR screening in order to clone cold-adapted lytic enzymes with degenerate primers deduced from conserved sequences of muramidases from Gram-negative bacteria. A Basic Local Alignment Search Tool (BLAST) sequence comparison of the PCR products revealed highest similarity to the lytic transglycosylase C of the bacteria Pasteurella multocida, and 16S rRNA sequencing indicated highest similarity to the species Shewanella frigidimarina [152]. 3.1.3 Transferases The nitrate/nitrite permease (nrtP) gene of the marine cyanobacterium Synechococcus sp. was described and characterized by Sakamoto et al. NrtP is a member of the major facilitator superfamily and is unrelated to the ATPbinding cassette-type nitrate transporters that have been described for freshwater strains of cyanobacteria. The discovery of a nitrate/nitrite permease in Synechococcus sp. suggested that significant differences in nutrient transporters may occur in marine and freshwater cyanobacteria [153]. A DNA polymerase gene of Cenarchaeum symbiosum was identified in the vicinity of the rRNA operon on a large genomic contig. Its deduced amino acid sequence is highly similar to those of the archaeal family B (α-type) DNA polymerases. It shared highest overall sequence similarity with the crenarchaeal DNA polymerases from the extreme thermophiles Sulfolobus acidocal-

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darius and Pyrodictium occultum [154]. The cDNA nucleotide sequence of the genome segment B encoding the VP1 protein, the putative RNA-dependent RNA polymerase (RdRp), was determined for five marine birnavirus (MABV) strains from different host or geographic origins and one infectious pancreatic necrosis virus (IPNV) by Zhang and Suzuki [155]. This is the only example of a study on marine viral enzymes. The phylogenetic diversity of the ribulose-1,5-bisphosphate carboxylase/ oxygenase (RuBisCO) large-subunit genes of deep sea microorganisms was analyzed in samples from the mid-Atlantic Ridge and various deep sea habitats around Japan including symbiont-bearing tissues of the vent mussel, Bathymodiolus sp., and the seep vestimentiferan tubeworm, Lamellibrachia sp. The RuBisCO sequences from the symbiont-bearing tissues showed a phylogenetic relationship with those from the ambient bacteria [156]. 3.2 Cloning and Gene Characterization The cloning and expression of enzymes are still considered a challenging job for marine biotechnologists. Nevertheless, cloning and characterization of genes encoding for carbohydrases, proteases, and antioxidant enzymes have been reported in recent years. In this section, the enzymes described are classified as hydrolases, oxidoreductases, and transferases. Apart from these reports, some other marine enzymes that have been cloned and expressed are presented in Table 3 at the end of this section. The gene encoding an extracellular chi from marine Alteromonas sp. was cloned in E. coli JM109 using pUC18 [157]. One of the chi genes (chiC) of Alteromonas sp. was cloned, and the nucleotide sequence was determined by Tsujibo et al. [158]. Three chitinolytic clones were isolated from a genomic DNA library of Vibrio sp., a psychrotolerant bacterium from the Antarctic Ocean. The chiA of the isolate was overexpressed in E. coli BL21(DE3) [159]. Arthrobacter sp. isolated from the sea bottom along the Antarctic ice shell secretes two major chitinases, chiA and chiB (Arthrobacter chitinase ArChiA and ArChiB), in response to chitin induction. A single chromosomal DNA fragment containing the genes coding for both chitinases was cloned and sequenced in E. coli by Lonhienne et al. [160]. Alteromonas sp. secretes chiA, chiB, and chiC in the presence of chitin. A gene cluster involved in the chitinolytic system of the strain was cloned and sequenced upstream of and including the chiA gene. The gene cluster consisted of three different open reading frames organized in the order chiD, cellobiose phosphorylase (cbp)1, and chiA [161]. The chiB secreted by a marine Alteromonas sp. was purified, and the corresponding gene was also cloned and sequenced by Orikoshi et al. [162]. The gene encoding N-acetyl-β-glucosaminidase (cht) from the marine bacterium Alteromonas sp. was cloned into pUC18 in E. coli JM109, sequenced and designated as cht60 [163]. The manA gene encoding an extracellular β-1,4-mannanase of

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Table 3 Examples of some cloned marine enzymes Enzyme class

Enzyme

Source

Reference

Hydrolase

Poly (3-hydroxy butyrate) depolymerase

Alcaligenes faecalis

[174]

β-1,3-xylanase

Vibrio sp., Alcaligenes sp.

[175, 176]

Uracil-DNA glycosylase

Marine Gram-positive psychrophile

[177]

β-lactamase

Vibrio harveyi

[178]

Cellulase

Marine mussel, Mytilus edulis

[179]

Sphingolipid ceramide N-deacylase

Marine bacterium, Shewanella alga

[180]

β-agarase

Marine Pseudomonas sp.

[181]

Protein C kinase

Marine sponge, Geodia cydonium

[182]

Aspartate transcarbamylase

Deep-sea hyperthermophilic [183] archaeon Pyrococcus abyssi

ATP sulfurylase

Hydrothermal vent tubeworm Riftia pachyptila

[184]

Polyketide synthase

Streptomyces maritimus

[185]

Luciferase

Marine dinoflagellate, Gonyaulax polyedra Methylosulfonomonas methylovora

[186]

Alanine dehydrogenase

Marine psychrophilic bacterium

[188]

NADH-quinone reductase

Vibrio alginolyticus

[189]

Carbamoyl phosphate synthetase

Pyrococcus abyssi

[190]

Glutamine synthetase

Shewanella violacea

[191]

Transferase

Oxidoreductase

Sulfonic acid monooxygenase

Ligase

[187]

a marine bacterium, Vibrio sp., was cloned and sequenced [164]. Screening of expression libraries identified mannanase-positive clones in Rhodothermus marimus, a thermophilic bacterium isolated from marine hot springs [165]. Characterization in Thermotoga neapolitana of a catabolic gene cluster encoding two glycosyl hydrolases, 1,4-β-D-glucan glucohydrolase and cellobiose phosphorylase, was reported by Yernool et al. [166]. To achieve a better un-

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derstanding of the molecular mechanisms underlying amylase expression, Ma et al. cloned and sequenced a 318-base pair (bp) fragment of amylase cDNA in developing sea bass (Lates calcarifer). Based on this sequence, a real-time reverse transcriptase PCR technique to monitor the changes in the messenger ribonucleic acid (mRNA) levels in the larvae was developed. A correlation between enzymatic activity and mRNA level of amylase could be demonstrated during the early development of sea bass larvae. This suggests that the changes in amylase are controlled at least at the transcriptional level during early larval development of sea bass [167]. A nuclear gene from the red alga Gracilaria gracilis was cloned by Lluisma and Ragan that encodes a homolog of starchbranching enzymes (SBEs) [168]. The complete cDNA coding for cathepsin L (hydrolase) was identified and characterized in the sponge Geodia cydonium by Krasko et al. The deduced amino acid sequence contains 322 residues, has a molecular weight of 36 085 Da, and shows the characteristic signatures known from other cathepsins of the L subfamily [95]. The gene encoding an extracellular alkaline metalloprotease was cloned, and its nucleotide sequence was analyzed and showed significant similarity to metalloproteases classified in the thermolysin family [169]. The encoding region of the gene of copper-zinc superoxide dismutase (oxidoreductase) has been cloned from several strains of marine yeast belonging to the genus Debaryomyces through genomic DNA-PCR amplification [170]. The encoding region of the superoxide dismutase enzyme gene of Udeniomyces puniceus was cloned from three species of pigmented marine yeast through genomic DNA-PCR amplification. For U. puniceus, the cloned nucleotide sequence contains all necessary information to produce a functional protein, which correlates with activity detected in cell homogenates [171]. Lluisma and Ragan cloned and sequenced a gene from G. gracilis that encodes a key enzyme of D-galactose metabolism, galactose-1-phosphate uridylyltransferase (GALT) [172]. A gene encoding an ADP-phosphofructokinase homolog has been identified in the hyperthermophilic archaeon Methanococcus jannaschii, which encodes a protein of 462 amino acids with a molecular weight of 53 361 Da. The gene was overexpressed in E. coli, and the produced enzyme was purified and characterized. The enzyme surprisingly showed high ADP-dependent activities for both glucokinase and phosphofructokinase [173].

4 Bioprocessing The area in which marine biotechnology in general and marine bioprocess engineering in particular have the greatest potential is in the design and optimization of bioreactors for marine metabolite production. A variety of bioreactor designs have been implemented, with varying degrees of success. The

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opportunity to produce new, bioactive structural analogs of known compounds via manipulation of culture conditions presents marine biotechnologists with a unique challenge for new bioproduct discovery. Innovations in media development, bioreactor design, and transgenic production, coupled with efficient downstream processing and product recovery, will be necessary to meet the needs of both discovery and bulk production of novel marine bioproducts [192]. With the discovery of and increased research into extremophiles, the conventional restrictions of low to moderate bioreactor temperatures and pressures faced by engineers may be circumvented [193]. Running bioprocess systems using marine hyperthermophiles poses interesting challenges such as bioreactor design and manipulation of their products, high temperature bioreactor operation, and corrosion of materials. Research in these areas examines the biotechnological potential of marine extremophiles from a biochemical engineering perspective [194]. Marine microorganisms often require special culture conditions such as high hydrostatic pressure in the case of deep sea bacteria and an optimized production medium for increased enzyme yield. They may also require an entirely different kind of complex nutrient in the production medium, which may be closer to the type of complex substances they are familiar with, unlike traditional sources such as soybean meal, corn steep liquor or molasses, or a chemically defined medium with known inducers [195]. A halotolerant strain of Bacillus licheniformis produced high protease activity during the early stationary phase of growth. The use of seawater in the production medium enhanced the production of this activity by 150% [196]. A marine bacterium V. harveyi was adapted to grow and produce extracellular proteases in a seawater/Zobellbased medium, supplemented with skim milk under different hydrodynamic conditions. The addition of skim milk to the Zobell medium enhanced the extracellular enzyme production fivefold. Specific growth rate increased as a consequence of increasing agitation rates [197]. Keerthi et al. reported Beauveria sp. isolated from marine sediment also produced extracellular L-glutaminase. Maximal L-glutaminase yield was obtained in a medium supplemented with yeast extract and sorbitol, sodium chloride, and methionine. This enzyme was inducible and growth associated [198]. Pseudomonas pseudomelli produced maximal chitinase activities during the late exponential and stationary phase under submerged fermentation. Manganese significantly enhanced chitinase production [199]. Immobilized cell technology has received the attention of marine biotechnologists. Cells from a marine bacterium, Teredinobacter turnirae, were immobilized in calcium alginate beads and used for alkaline protease production. There was no significant difference in the maximum protease activity between the three bead sizes used. A drastic fall in protease production was observed when the beads were treated with glutaraldehyde. The beads were used for eight successive fermentation batches, each lasting 72 h. It was also observed by Elibol

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and Moreira that there was an ∼ 3.5-fold increase in volumetric productivity of protease after the fourth cycle [200]. A marine Pseudomonas sp. immobilized by Ca-alginate gel entrapment was used for the production of extracellular Lglutaminase under repeated batch process and continuous process employing a packed bed reactor by Kumar and Chandrasekaran. In general, the volumetric productivity increased with increased dilution rate and substrate concentrations, and the substrate conversion efficiency declined [201]. Recently some investigations have been reported on the applications of solid state fermentation (SSF) in marine bioprocessing. Prawn waste, a chitinous solid waste of the shellfish processing industry, was used as a substrate for chitinase production by the marine fungus Beauveria bassiana in a SSF culture. The process parameters influencing SSF were optimized. The results indicate the scope for the use of shellfish processing (prawn) waste for the industrial production of chitinase by using SSF [202]. A chitinolytic fungus B. bassiana was isolated from marine sediment, and significant process parameters influencing chitinase production in SSF using wheat bran were optimized [203]. Process parameters influencing L-glutaminase production by marine V. costicola in SSF using polystyrene as an inert support were optimized [204]. Extracellular Lglutaminase production by Beauveria sp., isolated from marine sediment, was observed during SSF using polystyrene as an inert support by Sabu et al. [205]. Results indicate the scope for the production of salt-tolerant L-glutaminase using this marine fungus.

5 Conclusion In the past decade, of the plentiful reports on enzymes from novel and exotic sources few have reached the stage of commercial production. The problem lies in providing the enzyme producers with the proper environmental conditions of their ecological niches. Sustained production of the bioactive molecules by novel molecular methods of gene cloning and expression and innovative bioreactor designs like the so-called “niche-mimic” bioreactors [206] should play a pivotal role. Marine enzyme biotechnology will be the focus of the industry in the future. With mutual respect for each other’s commercial interests and intellectual property rights, the biodiverse but resource-poor developing world and the wealthy but bioresource-scarce developed world should join hands to unravel the secrets of this unopened research treasure chest—the world’s oceans. Acknowledgements Financial support through an ICMR Senior Research Fellowship to Malay Saha and research grants (AICTE and DST, Government of India) to Joydeep Mukherjee are thankfully acknowledged.

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References 1. Ireland CM, Copp BR, Foster MD, McDonald LA, Radisky DC, Swersey JC (1993) Marine Biotechnology, Vol 1. In: Attaway DH, Zaborsky OR (eds) Pharmaceutical and Bioactive Natural Products. Plenum, New York, pp 1–43 2. Zilinskas RA, Colwell RR, Lipton DW, Hill RT (1995) The global challenge of marine biotechnology: a status report on the United States, Japan, Australia and Norway Maryland Sea Grant, College Park (MD), p 372 3. Colwell RR (2002) Biotechnol Adv 20:215 4. Grace ES (1997) Biotechnology in Seas and Trees, 6. In: Biotechnology Unzipped: Promises and Realities, Natl Academies Press, Washington, DC, p 170 5. Wells W (1999) http://www.accessexcellence.org/ 6. Deming JW (2002) Curr Opin Microbiol 5:301 7. Deming JW (1998) Curr Opin Biotechnol 9:283 8. Horikoshi K(1998) Curr Opin Microbiol 1:291 9. Michels PC, Clark DS (1997) Appl Environ Microbiol 63:3985 10. Dib R, Chobert JM, Dalgalarrondo M, Barbier G, Haertle T (1998) FEBS Lett 431:279 11. Chavez CP, Sako Y, Uchida A (1999) Extremophiles 3:3 12. Francesco C (2002) International Symposium on Natural Products from Marine Microorganisms. Greifswald, Germany 13. Legin E, Ladrat C, Godfroy A, Barbier G, Duchiron F (1997) Comptes Rendus de l’Académie des Sciences - Series III - Sciences de la Vie 320:893 14. Brown SH, Kelly RM (1993) Appl Environ Microbiol 59:2614 15. Gantelet H, Duchiron F (1998) Appl Microbiol Biotechnol 49:770 16. Huber R, Stöhr J, Hohenhaus S, Rachel R, Burggraf S, Jannasch HW, Karl O, Stetter KO (1995) Arch Microbiol 164:255 17. Nishihara H, Miyashita Y, Aoyama K, Kodama T, Igarashi Y, Takamura Y (1997) Biochem Biophys Res Commun 232:766 18. Mukhopadhyay B, Patel VJ, Wolfe RS (2000) Arch Microbiol 174:406 19. Roy R, Adams MWW (2002) J Bacteriol 184:6952 20. Soda K, Kazuoka T, Fukui Y, Oikawa T (2002) International Symposium on Natural products from marine microorganisms, Greifswald, Germany 21. Irwin JA, Gudmundsson HM, Marteinsson VT, Hreggvidsson GO, Lanzetti AJ, Alfredsson GA, Engel PC (2001) Extremophiles 5:199 22. Irwin JA, Alfredsson GA, Lanzetti AJ, Gudmundsson HM, Engel PC (2001) FEMS Microbiol Lett 201:285 23. Kazuoka T, Masuda Y, Oikawa T, Soda K (2003) J Biochem (Tokyo) 133:51 24. Watanabe S, Takada Y, Fukunaga N (2001) Biosci Biotechnol Biochem 65:1095 25. Iyo AH, Forsberg CW (1999) Appl Environ Microbiol 65:995 26. Kammel U, Mennenga B, Hanschke R, Helmke E, Heinrich HW, Schweder T (2002) International Symposium on Natural products from marine microorganisms, Greifswald, Germany 27. Feller G, Zekhnini Z, Lamotte-Brasseur J, Gerday C (1997) Eur J Biochem 244:186 28. Konisky J, Michels PC, Clark DS (1995) Appl Environ Microbiol 61:2762 29. Qureshi MH, Kato C, Horikoshi K (1998) Extremophiles 2:93 30. Greene RV, Griffin HL, Cotta MA (1995) Tektran, Agricultural Research Service, United States Department of Agriculture, Washington, DC 31. Minic Z, Simon V, Penverne B, Gaill F, Hervé G (2001) J Biol Chem 276:23777 32. Minic Z, Pastra-Landis S, Gaill F, Hervé G (2003) J Biol Chem 278:40527 33. Cavanaugh CM, Wirsen CO, Jannasch HW (1992) Appl Environ Microbiol 58:3799

214

G. Debashish et al.

34. Turkiewicz M, Kur J, Bialkowska A, Cieslinski H, Kalinowska H, Bielecki S (2003) Biomol Eng 20:317 35. Mohapatra BR (1997) Indian J Mar Sci 26:292 36. Mohapatra BR, Banerjee UC, Bapuji M (1998) J Biotechnol 60:113 37. Mohapatra BR, Bapuji M (1997) Lett Appl Microbiol 25:393 38. Mohapatra BR, Bapuji M (1998) J Appl Microbiol 84:393 39. Turkiewicz M, Gromek E, Kalinowska H, Zielinska M (1999) J Biotechnol 70:53 40. Dunlap PV, Callahan SM (1993) J Bacteriol 175:4615 41. Callahan SM, Cornell NW, Dunlap PV (1995) J Biol Chem 270:17627 42. Dhevendaran K, Annie K (1998) Indian J Mar Sci 28:335 43. Mohapatra BR, Sani RK, Banerjee UC (1996) Lett Appl Microbiol 21:380 44. Mohapatra BR, Bapuji M, Banerjee UC (1998) Cytobios 92:165 45. Sawabe T, Ohtsuka M, Ezura Y (1997) Carbohydr Res 304:69 46. Biotechnology for the 21st century: new horizons 5. Opportunities in marine biotechnology and aquaculture (1995) Reports prepared by Biotechnology Research Subcommittee, Committee on Fundamental Science and National Science and Technology Council, USA 47. Shibata M, Takahashi S, Sato R, Oda K (1997) Biosci Biotechnol Biochem 61:710 48. Obata H, Sugiyama A, Kawahara H, Muramatsu T (1997) Biosci Biotechnol Biochem 61:1102 49. Miyamoto K, Nukui E, Hirose M, Nagai F, Sato T, Inamori Y, Tsujibo H (2002) Appl Environ Microbiol 68:5563 50. Rath J, Herndl GJ (1994) Appl Environ Microbiol 60:807 51. Arrieta JM, Herndl GJ (2001) Appl Environ Microbiol 67:4896 52. Arellano-Carbajal F, Olmos-Soto J (2002) World J Microbiol Biotechnol 18:791 53. Burtseva YV, Verigina NS, Sova VV, Pivkin MV, Zvyagintseva TN (2003) Mar Biotechnol 5:349 54. Arnosti C, Repeta DJ (1994) Appl Environ Microbiol 60:840 55. Tamaru Y, Araki T, Amagoi H, Mori H, Morishita T (1995) Appl Environ Microbiol 61:4454 56. Leon O, Quintana L, Peruzzo G, Slebe JC (1992) Appl Environ Microbiol 58:4060 57. Sugano Y, Terada I, Arita M, Noma M, Matsumoto T (1993) Appl Environ Microbiol 59:1549 58. Sugano Y, Kodama H, Terada I, Yamazaki Y, Noma M (1994) J Bacteriol 176:6812 59. Vera J, Alvarez R, Murano E, Slebe JC, Leon O (1998) Appl Environ Microbiol 64:4378 60. Kim BJ, Kim HJ, Ha SD, Hwang SH, Byun DS, Lee TH, Kong JY (1999) Biotechnol Lett 21:1011 61. Whitehead LA, Stosz SK, Weiner RM (2001) Cytobios 106:99 62. Tsujibo H, Yoshida Y, Miyamoto K, Imada C, Okami Y, Inamori Y (1992) Can J Microbiol 38:891 63. Svitil AL, Chadhain SMN, Moore JA, Kirchman DL (1997) Appl Environ Microbiol 63:408 64. Vaidya SY, Vala AK, Dube HC (2000) Indian J Mar Sci 29:336 65. Araki T, Tani S, Maeda K, Hashikawa S, Nakagawa H, Morishita T (1999) Biosci Biotechnol Biochem 63:2017 66. D’Souza MBD, Nair S, Chandramohan D (2000) Indian J Mar Sci 29:48 ´ S, A ´ sgeirsson B (2000) Enzyme Microb Technol 27:66 67. Hauksson JB, Andrésson O 68. Brandão PFB, Bull AT (2003) Antonie Van Leeuwenhoek 84:89 69. Colonna S, Gaggero N, Richelmi C, Pasta P (1999) Trends Biotechnol 17:163

Marine Enzymes

215

70. Raghukumar C, D’Souza TM, Thorn RG, Reddy CA (1999) Appl Environ Microbiol 65:2103 71. Francis CA, Co EM, Tebo BM (2001) Appl Environ Microbiol 67:4024 72. Yang Q, Yang M, Pritsch K, Yediler A, Hagn A, Schloter M, Kettrup A (2003) Biotechnol Lett 25:709 73. Chadd HE, Newman J, Mann NH, Carr NG (1996) FEMS Microbiol Lett 138:161 74. Hernandez-Saavedra NY, Ochoa JL (1999) Yeast 15:657 75. Tsugawa W, Horiuchi S, Tanaka M, Wake H, Sode K (1996) Appl Biochem Biotechnol 56:301 76. Tsugawa W, Ogasawara N, Sode K (1998) Enzyme Microb Technol 22:269 77. Besson S, Carneiro C, Moura JJG, Moura I, Fauque G (1995) Anaerobe 1:219 78. De Souza MP, Yoch DC (1995) Appl Environ Microbiol 61:21 79. De Souza MP, Yoch DC (1995) Appl Environ Microbiol 61:3986 80. Jansen M, Hansen TA (2000) J Sea Res 43:225 81. Ansede JH, Pellechia PJ, Yoch DC (2001) Appl Environ Microbiol 67:3134 82. Sakai T, Kimura H, Kojima K, Shimanaka K, Ikai K, Kato I (2003) Mar Biotechnol 5:380 83. Carpenter EJ, Bergman B, Dawson R, Siddiqui PJ, Soderback E, Capone DG (1992) Appl Environ Microbiol 58:3122 84. Alaoui SE, Diez J, Humanes L, Toribio F, Partensky F, García-Fernández JM (2001) Appl Environ Microbiol 67:2202 85. Stickforth P, Steiger S, Hess WR, Sandmann G (2003) Arch Microbiol 179:409 86. Laroche M, Ulber R (2002) International Symposium on Natural Products from Marine Microorganisms, Greifswald, Germany 87. Kita K, Ishimaru K, Troika M, Yanase H, Kato N (1995) Appl Environ Microbiol 61:1727 88. Simpson H, Denis M, Malatesta F (1997) Biosci Rep 17:343 89. Barbeyron T, Potin P, Richard C, Collin O, Kloareg B (1995) Microbiology 141:2897 90. Ghosh D (2004) PhD Thesis, Jadavpur University, Kolkata, India 91. Nilsen IW, Overbo K, Sandsdalen E, Sandaker E, Sletten K, Myrnes B (1999) FEBS Lett 464:153 92. Gregersen F (2000) Norwegian Institute of Fisheries and Aquaculture (NIFA) Fiskeriforskning Info No. 3 July 2000 93. Nilsen IW, Myrnes B (2001) Gene 269:27 94. Arreguín R, Arreguín B, Soriano-García M, Hernández-Arana A, Rodríguez-Romero A (1993) FEBS Lett 320:235 95. Krasko A, Gamulin V, Seack J, Steffen R, Schroder HC, Muller WE (1997) Mol Mar Biol Biotechnol 6:296 96. Sivakumar P, Sampath P, Chandrakasan G (1999) Comp Biochem Physiol B Biochem Mol Biol 123:273 97. Munilla-Morán R, Saborido-Rey F (1996) Comp Biochem Physiol B Biochem Mol Biol 113:827 98. Mora P, Fournier D, Narbonne J (1999) Comp Biochem Physiol C Pharmacol Toxicol Endocrinol 122:353 99. Lushchak VI, Smirnova YD, Storey KB (1998) Comp Biochem Physiol B Biochem Mol Biol 119:611 100. Lucu C, Devescovi M, Skaramuca B, Kozul V (2000) J Exp Mar Biol Ecol 246:163 101. Frost GI, Csoka T, Stern R (1996) Trends Glycosci Glycotechnol 8:419 102. Reintamm T, Lopp A, Kuusksalu A, Pehk T, Kelve M (2003) Eur J Biochem 270:4122 103. Shigeta S, Suzuki O, Aki Y, Kawamoto S, Ono K (2000) J Biosci Bioeng 89:84 104. Saborowski R, Buchholz F (2002) Mar Biol 140:557

216

G. Debashish et al.

105. Noguchi K, Ishikawa K, Yokoyama K, Ohtsuka T, Nio N, Suzuki E (2001) J Biol Chem 276:12055 106. Angelucci S, Sacchetta P, Moio P, Melino S, Petruzzelli R, Gervasi PG, Di Ilio C (2000) Arch Biochem Biophys 373:435 107. MacDonald JA, Storey KB (1999) Mar Biol 133:193 108. Chen YP, Woodin SA, Lincoln DE, Lovell CR (1996) J Biol Chem 271:4609 109. Birmelin C, Pipe RK, Goldfarb PS, Livingstone DR (1999) Mar Biol 135:65 110. Novi S, Pretti C, Cognetti AM, Longo V, Marchetti S, Gervasi PG (1998) Aquat Toxicol 41:63 111. Rengasamy A, Munusamy A, Periasamy M (1997) Develop Comp Immunol 21:1 112. Kazeto Y, Adachi S, Yamauchi K (2001) Gen Comp Endocrinol 122:109 113. Ali SA, Alam JM, Stoeva S, Schütz J, Abbasi A, Zaidi ZH, Voelter W (1999) Toxicon 37:1505 114. Liu DL, Walden CC (1997) Fish Res Bd Canada 27:1141 115. Moore RM, Webb M, Tokarczyk R, Wever R (1996) J Geophys Res 101:899 116. Ohsawa N, Ogata Y, Okada N, Itoh N (2001) Phytochemistry 58:683 117. Martinez JS, Carroll GL, Tschirret-Guth RA, Altenhoff G, Little RD, Butler A (2001) J Am Chem Soc 123:3289 118. Touchette BW, Burkholder J (2001) Plant Physiol Biochem 39:583 119. Ueno Y, Kurano N, Miyachi S (1999) FEBS Lett 443:144 120. Morishita H, Ohashi S, Oku T, Nakajima Y, Kojima S, Ryufuku M, Nakamura H, Ohmiya Y (2002) Photochem Photobiol 75:311 121. Deng T, Roenneberg T (2002) Naturwissenschaften 89:171 122. Matsubara K, Hori K, Matsuura Y, Miyazawa K (2000) Comp Biochem Physiol B Biochem Mol Biol 125:137 123. Gupta A, Vijayaraghavan MR, Gupta R (1998) Phytochemistry 49:1875 124. Muramatsu Y, Harada A, Ohwaki Y, Kasahara Y, Takagi S, Fukuhara T (2002) Plant Cell Physiol 43:1137 125. Fan C, Glibert PM, Alexander J, Lomas MW (2003) Mar Biol 142:949 126. Satoh D, Hiraoka Y, Colman B, Matsuda Y (2001) Plant Physiol 126:1459 127. Wise ML, Rorrer GL, Polzin JJ, Croteau R (2002) Arch Biochem Biophys 400:125 128. Maurin C, Gal YL (1997) Plant Sci 122:61 129. Pichard SL, Campbell L, Paul JH (1997) Appl Environ Microbiol 63:3600 130. Zheng W, Wise ML, Wyrick A, Metz JG, Yuan L, Gerwick WH (2002) Arch Biochem Biophys 401:11 131. Krasko A, Lorenz B, Bate R, Schröder HC, Müller IM, Müller WE (2000) Eur J Biochem 267:4878 132. Pace NR (1997) Science 276:734 133. Short JM (1997) Nat Biotechnol 15:1322 134. DeLong EF (2002) Curr Opin Microbiol 5:520 135. Saiki RK, Gelfand DH, Stoffel S, Scharf SJ, Higuchi R, Horn GT, Mullis KB, Erlich HA (1988) Science 239:487 136. Giovannoni SJ, Britschgi TB, Moyer CL, Field KG (1990) Nature 345:60 137. Braker G, Zhou J, Wu L, Devol AH, Tiedje JM (2000) Appl Environ Microbiol 66:2096 138. Grüntzig V, Nold SC, Zhou J, Tiedje JM (2001) Appl Environ Microbiol 67:760 139. Scala DJ, Kerkhof LJ (1999) Appl Environ Microbiol 65:1681 140. Zehr JP, Mellon MT, Zani S (1998) Appl Environ Microbiol 64:3444 141. Mehta MP, Butterfield DA, Baross JA (2003) Appl Environ Microbiol 69:960 142. Bano N, Hollibaugh JT (2000) Appl Environ Microbiol 66:1960 143. Nold SC, Zhou J, Devol AH, Tiedje JM (2000) Appl Environ Microbiol 66:4532

Marine Enzymes

217

144. Buchan A, Neidle EL, Moran MA (2001) Appl Environ Microbiol 67:5801 145. Hedlund BP, Geiselbrecht AD, Bair TJ, Staley JT (1999) Appl Environ Microbiol 65:251 146. Geiselbrecht AD, Hedlund BP, Tichi MA, Staley JT (1998) Appl Environ Microbiol 64:4703 147. Cottrell MT, Wood DN, Yu L, Kirchman DL (2000) Appl Environ Microbiol 66:1195 148. Svitil AL, Kirchman DL (1998) Microbiology 144:1299 149. Cottrell MT, Moore JA, Kirchman DL (1999) Appl Environ Microbiol 65:2553 150. Collier JL, Brahamsha B, Palenik B (1999) Microbiology145:447 151. Wiens M, Koziol C, Batel R, Müller WE (1999) Mar Biotechnol 1:191 152. Borriss M, Hanschke R, Helmke E, Heinrich H, Schweder T (2002) International Symposium on Natural Products from Marine Microorganisms, Greifswald, Germany 153. Sakamoto T, Inoue-Sakamoto K, Bryant DA (1999) J Bacteriol 181:7363 154. Schleper C, Swanson RV, Mathur EJ, DeLong EF (1997) J Bacteriol 179:7803 155. Zhang CX, Suzuki S (2003) Arch Virol 148:745 156. Elsaied H, Naganuma T (2001) Appl Environ Microbiol 67:1751 157. Tsujibo H, Orikoshi H, Tanno H, Fujimoto K, Miyamoto K, Imada C, Okami Y, Inamori Y (1993) J Bacteriol 175:176 158. Tsujibo H, Orikoshi H, Shiotani K, Hayashi M, Umeda J, Miyamoto K, Imada C, Okami Y, Inamori Y (1998) Appl Environ Microbiol 64:472 159. Bendt A, Hüller H, Kammel U, Helmke E, Schweder T (2001) Extremophiles 5:119 160. Lonhienne T, Mavromatis K, Vorgias CE, Buchon L, Gerday C, Bouriotis V (2001) J Bacteriol 183:1773 161. Tsujibo H, Orikoshi H, Baba N, Miyahara M, Miyamoto K, Yasuda M, Inamori Y (2002) Appl Environ Microbiol 68:263 162. Orikoshi H, Baba N, Nakayama S, Kashu H, Miyamoto K, Yasuda M, Inamori Y, Tsujibo H (2003) J Bacteriol 185:1153 163. Tsujibo H, Fujimoto K, Tanno H, Miyamoto K, Imada C, Okami Y, Inamori Y (1994) Gene 146:111 164. Tamaru Y, Arakia T, Morishita T, Kimura T, Sakka K, Ohmiya K (1997) J Ferment Bioeng 83:201 165. Politz O, Krah M, Thomsen KK, Borriss R (2000) Appl Microbiol Biotechnol 53:715 166. Yernool DA, McCarthy JK, Eveleigh DE, Bok J (2000) J Bacteriol 182:5172 167. Ma P, Sivaloganathan B, Reddy PK, Chan WK, Lam TJ (2001) Mar Biotechnol 3:463 168. Lluisma AO, Ragan MA (1998) Curr Genet 34:105 169. Miyamoto K, Tsujibo H, Nukui E, Itoh H, Kaidzu Y, Inamori Y (2002) Biosci Biotechnol Biochem 66:416 170. Hernandez-Saavedra NY, Romero-Geraldo R (2001) Yeast 18:1227 171. Hernandez-Saavedra NY (2003) Yeast 20:479 172. Lluisma AO, Ragan MA (1998) Curr Genet 34:112 173. Ohshima T, Sakuraba H (2002) International Symposium on Natural Products from Marine Microorganisms, Greifswald, Germany 174. Kita K, Mashiba S, Nagita M, Ishimaru K, Okamoto K, Yanase H, Kato N (1997) Biochim Biophys Acta 1352:113 175. Araki T, Hashikawa S, Morishita T (2000) Appl Environ Microbiol 66:1741 176. Okazaki F, Tamaru Y, Hashikawa S, Li Y, Araki T (2002) J Bacteriol 184:2399 177. Jaeger S, Schmuck R, Sobek H (2000) Extremophiles 4:115 178. Teo JWP, Suwanto A, Poh CL (2000) Antimicrob Agents Chemother 44:1309 179. Xu B, Janson JC, Sellos D (2001) Eur J Biochem 268:3718 180. Furusato M, Sueyoshi N, Mitsutake S, Sakaguchi K, Kita K, Okino N, Ichinose S, Omori A, Ito M (2002) J Biol Chem 277:17300

218

G. Debashish et al.

181. Kang NY, Choi YL, Cho YS, Kim BK, Jeon BS, Cha JY, Kim CH, Lee YC (2003) Biotechnol Lett 25:1165 182. Seack J, Kruse M, Isabel M, Müller IM, Müller WE (1999) Biochim Biophys Acta 1444:241 183. Purcarea C, Herve G, Ladjimi MM, Cunin R (1997) J Bacteriol 179:4143 184. Laue BE, Nelson DC (1994) J Bacteriol 176:3723 185. Piel J, Hertweck C, Shipley PR, Hunt DM, Newman MS, Moore BS (2000) Chem Biol 7:943 186. Bae YM, Hastings JW (1994) Biochim Biophys Acta 1219:449 187. D’Marco P, Moradas-Ferreira P, Higgins TP, McDonald I, Kenna EM, Murrell JC (1999) J Bacteriol 181:2244 188. Irwin JA, Lynch SV, Coughlan S, Baker PJ, Gudmundsson HM, Alfredsson GA, Rice DW, Engel PC (2003) Extremophiles 7:135 189. Nakayama Y, Hayashi M, Unemoto T (1998) FEBS Lett 422:240 190. Purcarea C, Hervé G, Cunin R, Evans DR (2001) Extremophiles 5:229 191. Ikegami A, Nakasone K, Kato C, Nakamura Y, Yoshikawa I, Usami R, Horikoshi K (2000) FEMS Microbiol Lett 192:91 192. Pomponi SA (1998) The Potential for the Marine Biotechnology Industry. Harbor Branch Oceanographic Institution, Florida; Trends and Future Challenges for US National Ocean and Coastal Policy, vol 101 193. Wright PC, Stevenson C, McEvoy E, Burgess JG (1999) J Biotechnol 70:343 194. Bustard M, Burgess JG, Meeyoo V, Wright PC (2000) J Chem Technol Biotechnol 75:1095 195. Chandrasekaran M (1997) J Mar Biotechnol 5:86 196. Manachini PL, Fortina MG (1998) Biotechnol Lett 20:565 197. César EB, Facundo MR (2003) World J Microbiol Biotechnol 19:129 198. Keerthi TR, Suresh PV, Sabu A, Kumar SR, Chandrasekaran M (1999) World J Microbiol Biotechnol 15:751 199. Suresh PV, Sindhu R, Chandrasekaran M (1995) National symposium on relevance of biotechnology in industry, Cochin, India 200. Elibol M, Moreira AR (2003) Process Biochem 38:1445 201. Kumar SR, Chandrasekaran M (2003) Process Biochem 38:1431 202. Suresh PV, Chandrasekaran M (1998) World J Microbiol Biotechnol 14:655 203. Suresh PV, Chandrasekaran M (1999) Process Biochem 34:257 204. Prabhu GN, Chandrasekaran M (1997) Process Biochem 32:285 205. Sabu A, Keerthi TR, Kumar SR, Chandrasekaran M (2000) Process Biochem 35:705 206. Yan L, Boyd KG, Adams DR, Burgess JG (2003) Appl Environ Microbiol 69:3719

Adv Biochem Engin/Biotechnol (2005) 96: 219–262 DOI 10.1007/b135786 © Springer-Verlag Berlin Heidelberg 2005 Published online: 26 August 2005

Extreme Environments as a Resource for Microorganisms and Novel Biocatalysts Garabed Antranikian1 (u) · Constantinos E. Vorgias2 · Costanzo Bertoldo1 1 Institute

of Technical Microbiology, Technical University Hamburg-Harburg, Kasernenstraße 12, 21073 Hamburg, Germany [email protected] 2 Faculty of Biology, Department of Biochemistry and Molecular Biology, National and Kapodistrian University of Athens, Panepistimiopolis-Zographou, 157 84 Athens, Greece [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 2.1 2.2 2.3 2.4

Extreme environments as a resource of unique genes and biocatalysts Low-temperature-adapted microorganisms . . . . . . . . . . . . . . . . Microorganisms that grow at elevated temperatures . . . . . . . . . . . Life at extremes of pH . . . . . . . . . . . . . . . . . . . . . . . . . . . . High-salt-tolerant microorganisms . . . . . . . . . . . . . . . . . . . . .

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Cellulases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Xylan-degrading enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Pectin-degrading enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Chitin-degrading enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Starch-processing enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . Heat-stable amylases, glucoamylases and α-glucosidases . . . . . . . . . . . Thermoactive pullulanase and CGTase . . . . . . . . . . . . . . . . . . . .

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Proteolytic enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Lipases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Glucose isomerases, alcohol dehydrogenases and esterases . . . . . . . . .

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Amidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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DNA-processing enzymes . . . . . Polymerase chain reaction (PCR) DNA sequencing . . . . . . . . . . Ligase chain reaction . . . . . . .

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Abstract The steady increase in the number of newly isolated extremophilic microorganisms and the discovery of their enzymes by academic and industrial institutions underlines the enormous potential of extremophiles for application in future biotechnological processes. Enzymes from extremophilic microorganisms offer versatile tools for sustainable developments in a variety of industrial application as they show important environmental benefits due to their biodegradability, specific stability under extreme conditions, improved use of raw materials and decreased amount of waste products. Although major advances have been made in the last decade, our knowledge of the physiology, metabolism, enzymology and genetics of this fascinating group of extremophilic microorganisms and their related enzymes is still limited. In-depth information on the molecular properties of the enzymes and their genes, however, has to be obtained to analyze the structure and function of proteins that are catalytically active around the boiling and freezing points of water and extremes of pH. New techniques, such as genomics, metanogenomics, DNA evolution and gene shuffling, will lead to the production of enzymes that are highly specific for countless industrial applications. Due to the unusual properties of enzymes from extremophiles, they are expected to optimize already existing processes or even develop new sustainable technologies. Keywords Extremophiles · Stable biocatalysts · Thermophiles · Extremes of pH · Psychrophiles · Enantioselectivity

1 Introduction Extremophiles are unique microorganisms that are adapted to survive in ecological niches such as high or low temperatures, extremes of pH, high salt concentrations and high pressure. Accordingly biological systems and enzymes can even function at temperatures between – 5 and 130 ◦ C, pH 0– 12, salt 3–35% and 1000 bar. The majority of the organisms that grow in these extreme environments belong to a group with distinct characteristics. Carl Woese named this group archaea, and postulated the archaea as the third domain of life on earth, different form bacteria and eukarya [1, 2]. A large number of these unique microorganisms have been isolated from marine environments (Table 1). In many cases microbial biocatalysts, especially of extremophiles, are superior to traditional catalysts, because they allow the performance of industrial processes even under harsh condition, under which conventional proteins are completely denatured. By virtue of their positive properties, stability, specificity, selectivity and efficiency, enzymes already occupy a prominent position in modern biotechnology. For many processes in the chemical and pharmaceutical industries, suitable microbial enzymes can be found that have the potential to optimize or even replace chemical processes. By using robust enzymes in biotechnical processes one is often able to better utilize raw materials, minimize pollutant emissions and reduce energy consumption while sim-

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Table 1 Some representatives of microorganisms living under extreme conditions Microbial life at various temperatures

Optimal growth (◦ C)

Phychrophiles Vibrio sp Micrococcus criophilus Arthrobacter glacialis Vibrio psychroerythreus Aquaspirillum articum

< 20 ◦ C < 20 ◦ C < 20 ◦ C < 20 ◦ C < 20 ◦ C

Moderate thermophiles (50–60 ◦ C) Bacillus acidocaldarius Bacillus stearothermophilus

50 55

Extreme thermophiles (60–80 ◦ C) Thermus aquaticus Thermoanaerobacter ethanolicus Clostridium thermusolfurogenes Fervidobacterium pennivorans

70 65 60 75

Hyperthermophiles (80–110 ◦ C) Thermotoga maritima Aquifex pyrophilus Archeoglobus fulgidus Methanopyrus kandleri Sulfolobus sulfataricus Thermococcus aggregans Pyrobaculum islandicum Pyrococcus furiosus Pyrodictium occultum Pyrolobus fumarii

90 85 83 88 88 88 100 100 105 106

ultaneously improving quality and purity of products, e.g. optically pure compounds. The additional benefits in performing industrial processes at high temperature include reduced risk of contamination, improved transfer rates, lower viscosity and higher solubility of substrates. The recent exciting results in the field of extremophile research, the high demands of the biotechnology industries for tailor-made novel biocatalysts and the simultaneous rapid development of new techniques will stimulate the development of innovative processes on the basis of biocatalyst from extremophiles.

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Table 1 (continued) Microbial life at various temperatures and pH

Optimal growth conditions Temperature pH

Acidophilic microorganisms Sarcina ventriculi Thiobacillus ferrooxidans Alyciclobacillus acidocaldarius Picrophilus oshimae Picrophilus torridus Thermoplasma acidophilum Sulfolobus acidocaldarius Acidianus infernus

37 37 55 60 60 60 75 75

4.0 2.5 2.0–6.0 0.7 0.7 2.0 2.5 2.0

60 85 85

9.5 9.0 9.0

Alkaliphilic microorganisms Anaerobranca gottschallki Thermococcus alcaliphilus Thermococcus acidoaminivorans

Range of NaCl concentration in M required for growth (optimum) Halophilic microorganisms Dunaliella spp. Clostridium halophilum Haloanaerobium praevalens Halobacterium denitrificans Haloferax vulcanii

0.3–5.0 0.15–6.0 (0.6) 0.8–4.3 (2.2) 1.5–4.5 (2.5) 1.0–3.0 (1.5)

2 Extreme environments as a resource of unique genes and biocatalysts Modern biotechnology has a steadily increasing demand for novel enzymes. The classical approach to the isolation of new or improved biocatalysts requires that the different microorganisms derived from an environmental sample be cultured on an appropriate growth medium and separated until individual colonies are isolated. The enzymes and the corresponding genes are then recovered from the identified microorganism. This value of the classical method is substantial but it fails, however, to represent the scope of microbial diversity in nature, since only a small proportion (1–3%) of viable microorganisms in a sample have to date been recovered by culturing techniques. To explore the diversity and the potential of microbial communities a method was developed to analyze DNA in environmental samples that bypasses classical cultivation techniques [3]. Collectively, the term metagenome indicates the genome of the total microbiota found in nature and contains vastly more genetic information than is contained in the cultivable subset. Hence, the

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construction of metagenomic libraries by direct extraction and cloning large fragments of DNA isolated directly from microbes in natural environments has represented an effective way of accessing the wealth of information of microbial mixed populations. This method circumvents the loss of major portions of the microbial communities in extreme environments, which derives from the different growth requirements of the different microbes. Several different laboratories have successfully isolated novel genes encoding different enzymes and secondary metabolites from microbial communities and their metagenomes without cultivation of the microbes [4]. The microbial niches studied were highly diverse and ranged from moderate environments, such as river soil [4], to extreme environments, such as the deep sea [5–9]. Studies have shown that the metagenomic approach offers a valuable pool for encountering novel genes encoding biotechnologically relevant gene products such as lipase [10] cellulases [11], amylases [12], chitinases [13, 14], and esterases [10]. Rondon et al. [15, 16] used a bacterial artificial chromosome (BAC) vector to express Bacillus cereus genomic DNA. The advantage of BAC vectors is that they maintain very large DNA inserts (greater than 100 kb) stably in E. coli, facilitating the cloning of large fragments of DNA. Their results demonstrated that expression of heterologous DNA from B. cereus in an E. coli BAC system was detectable at a reasonable frequency, validating the idea that the low-copy BAC vector (one to two per cell) could be used to express foreign DNA from foreign promoters in E. coli [15, 16]. However, a major difficulty associated with exploiting the metagenome directly from the environment is related to contamination of purified DNA with polyphenolic compounds that are copurified with the DNA. These compounds are difficult to remove, and it is well known that polyphenols also interfere with enzymatic modifications of isolated DNA [17, 18]. As a result, construction of environmentally derived DNA libraries with large inserts is hindered due to the poor quality of the isolated DNA. These known difficulties associated with the construction of libraries directly derived from environmental DNA samples forced researchers to isolate DNA from the metagenome of a microbial community after precultivation in the laboratory [19]. This technique, though it limits the biodiversity of laboratory enrichment cultures, has proven to be highly efficient for rapid isolation of large DNA fragments and for cloning of operons and genes with great biotechnological value. For example, Voget et al. [20] exploited the metagenome of an enrichment culture on agar plates for isolation of genes encoding a variety of different biocatalysts, including β-agarases, amylolytic enzymes, cellulases, lipases and many other enzymes with high biotechnological potential. Given the immense uncultivated and uncharacterized metabolic diversity in the environment, one would need to sequence a relatively restricted number of clones to discover fundamentally interesting sets of genes. However, this approach relies on the fortuitous expression of heterologous DNA by the library host strain. Although the speed and effectiveness of brute-force sequencing are constantly improving, it is not

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yet practical to assemble a complete bacterial genome from a metagenome. There is still a need for new functional genomic approaches that systematically yield information about many of the elements in a metagenomic library. If modern genomic techniques can be used to carry out more comprehensive surveys of metagenomic libraries, the understanding of natural genetic diversity would be greatly enhanced [21]. This technology has been established by various biotechnological industries such as Diversa in San Diego and B.r.a.i.n. in Germany. By applying a high-throughput system (HTS) extreme environments have been studied for the production of stable enzymes. 2.1 Low-temperature-adapted microorganisms The Earth’s biosphere is predominantly aqueous and cold. Nearly 70% is water and a high percentage of it seldom reaches temperatures above 5 ◦ C. The polar region provides a permanently cold environment that is surrounded by an aquatic belt of melting ice. Microorganisms able to grow at temperatures close to 0 ◦ C have developed various adaptation mechanisms to survive and function at low temperature. These microorganisms can be divided into two main groups: psychrophiles and psychrotolerants. Psychrophilic microorganisms grow at an optimum temperature of 15 ◦ C, with a maximum growth temperature at about 20 ◦ C and a minimum around 0 ◦ C. Psychrotolerant microorganisms generally do not grow at zero but do so at 3–5 ◦ C, and have optimum and maximum growth temperatures above 20 ◦ C but less than 30 ◦ C [22]. Most of the cold-adapted microorganisms have been characterized from Arctic and Antarctic seawater and, despite the harsh conditions, the density of bacterial cells in the Antarctic oceans is as high as the density reported in temperate waters (Table 1). Psychrophiles can be found in permanently cold environments such as the deep sea, glaciers, and mountain regions, in soils, in fresh or saline waters associated with cold-blooded animals such as fish or crustaceans. In general, cold-adapted enzymes have higher specific activity at low and moderate temperatures than that of their mesophilic counterparts, and are inactivated easily by a slight increase in temperature [23, 24]. 2.2 Microorganisms that grow at elevated temperatures Microorganisms capable of growing optimally at temperatures between 50 and 60 ◦ C are designated as moderate thermophiles (Table 1). Most of these microorganisms belong to many different taxonomic groups of eu- and prokaryotic microorganisms such as protozoa, fungi, algae, streptomycetes and cyanobacteria, which comprise mainly mesophilic species. It can be assumed that moderate thermophiles, which are closely related phylogenetically

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to mesophilic organisms, may be secondarily adapted to life in hot environments. Extreme thermophiles, which grow optimally between 60 and 80 ◦ C, are widely distributed among the genera Bacillus, Clostridium, Thermoanaerobacter, Thermus, Fervidobacterium, Thermotoga and Aquifex. The relative abundance of archaea and bacteria in high-temperature environments was, until recently, mainly studied by cultivation-based techniques. Because of the frequent isolation of archaea from these habitats, it was assumed that archaea dominate the high-temperature biotopes. Recently, the application of molecular-biological methods revealed that bacterial communities are also abundant in these environments. These results suggest that archaea may generally be of lower abundance in hot environments than could be assumed from cultivation-based experiments. However, the factors that allow bacteria to dominate in high-temperature habitats, that were once believed to be the realm of archaea, remain unknown [25, 26]. Microorganisms that are adapted to grow optimally at very high temperatures (80–108 ◦ C) have been isolated from high-temperature terrestrial and marine habitats. The most common biotopes are volcanically and geothermal-heated hydrothermal vent systems such as solfataric fields, neutral hot springs, and submarine hot vents. Submarine hydrothermal systems are situated at shallow and abyssal depth. They consist of hot fumaroles, springs, sediments, and deep-sea vents with temperatures of up to 400 ◦ C (“black smokers”) [27]. Because of their ability to convert volcanic gases and sulphur compounds at high temperatures, hyperthermophilic communities living in such hydrothermal vents are expected to play an important role in marine ecological, geochemical and volcanic processes [28]. Shallow as well as deep-sea hydrothermal systems harbour members of various genera including Pyrococcus, Pyrodictium, Igneococcus, Thermococcus, Methanococcus, Archaeoglobus and Thermotoga. So far, members of the genus Methanopyrus have been found only at greater depths, whereas Aquifex was isolated exclusively from shallow hydrothermal vents. Recently, interesting biotopes of extreme and hyperthermophiles were discovered in deep, geothermally heated oil reservoirs around 3500 m below the bed of the North Sea and the permafrost soil of North Alaska [29]. Interestingly, the majority of the hyperthermophiles isolated to date belong to the archaeal domain of life and no eukaryotic organism has been found that can grow at the boiling point of water. A 16S rDNA-based universal phylogenetic tree shows a tripartite division of the living world consisting of the domains Bacteria, Archaea and Eukarya [1, 30]. 2.3 Life at extremes of pH Solfataric fields are the most important biotopes of microorganisms that prefer to live under both thermophilic and acidic conditions. Solfataric soils con-

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sist of two different layers that can be easily distinguished by their characteristic colours: the upper, aerobic layer has an ochre colour due to the presence of ferric iron. The layer below, which is anaerobic, appears rather blackishblue owing to the presence of ferrous iron. Thermophilic acidophiles, belonging to the genera Sulfolobus [31, 32], Acidianus [33], Thermoplasma [34], and Picrophilus [35], with growth optima between 60 and 90 ◦ C and pH 0.7–5.0 are commonly found in the aerobic upper layer, whereas slightly acidophilic or neutrophilic anaerobes such as Thermoproteus tenax or Methanothermus fervidus can be isolated from the lower layer. Species of Thermoplasma (growth optima: pH 2.0 and 60 ◦ C) have been found in hot springs, solfataras and coal refuse piles [36]. Their closest known phylogenetic relatives, also found in solfataras, are species of the genus Picrophilus, which are so far the most extreme acidophiles, with growth close to pH 0. Picrophilus oshimae and P. torridus are both aerobic, heterotrophic archaea that grow optimally at 60 ◦ C and pH 0.7 and utilize various polymers such as starch and proteins as carbon source (Table 1) [37, 38]. Members of the genus Sulfolobus are strict aerobes growing either autotrophically, heterotrophically or facultative heterotrophically. During autotrophic growth, S0 , S2– and H2 are oxidized to sulphuric acid or water as end products. Sulfolobus metallicus [39–42] and S. brierley [43] are able to grow by oxidation of sulfidic ores. A dense biofilm of these microorganisms is responsible for the microbial ore leaching process, in which heavy-metal ions such as Fe2+ , Zn2+ and Cu2+ are solubilized. Other thermoacidophiles have been affiliated to the genera Metallosphaera (growth range: 50–80 ◦ C, pH 1–4.5) [44], Acidianus (growth range: 60–95 ◦ C, pH 1.5–5) [33] and Stygioglobus (growth range: 57–90 ◦ C, pH 1–5.5). The alkaliphiles that grow at high pH values are widely distributed throughout the world. They have been found in carbonate-rich springs and alkaline soils, where the pH can be around 10.0 or even higher, although the internal pH is maintained around 8.0. In such places, several species of cyanobacteria and Bacillus are normally abundant and provide organic matter for diverse groups of heterotrophs [45]. Alkaliphiles require alkaline environments and sodium ions not only for growth but also for sporulation and germination. Sodium-ion-dependent uptakes of nutrients have been reported in alkaliphiles. Many alkaliphiles require various nutrients for growth; few alkaliphilic Bacillus strains can grow in simple minimal media containing glycerol, glutamic acid, and citric acid [46]. In general, cultivation temperature is in the range of 20–55 ◦ C. Furthermore, many haloalkaliphiles isolated from alkaline hypersaline lakes can grow in alkaline media containing 20% NaCl. The soda lakes in the Rift Valley of Kenya and similar lakes found in a few other places on earth are highly alkaline with pH values between 11.0 or 12.0 and represent a typical habitat where alkaliphilic microorganisms can be isolated [47]. Thermophilic anaerobic spore-forming alkaliphiles, thermoalkaliphilic Clostridia, were isolated from sewage plants [48]. Very recently,

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two thermoalkaliphilic bacteria, Anaerobranca gottschalkii and Anaerobranca horikoshii have been isolated from Lake Bogoriae in Kenya and from Yellowstone National Park, respectively [49, 50] (Table 1). The new isolates represent a new line within the Clostridium/Bacillus subphylum. The two archaeal thermoalkaliphiles identified to date are Thermococcus alcaliphilus [51] and Thermococcus acidoaminivorans [52], both growing at 85 ◦ C and pH 9.0. 2.4 High-salt-tolerant microorganisms The halophiles comprise Bacteria and Archaea that grow optimally at NaCl concentrations above that of seawater (> 0.6 M NaCl). In general, halophilic microorganisms are classified as moderate halophiles if they can grow at salt concentrations between 0.85 and 1.7 M NaCl and as extreme halophiles if they require NaCl concentrations above 1.7 M for growth. Halophiles have been mainly isolated from saline lakes, such as the Great Salt Lake in Utah (salinity > 2.6 M) and from evaporated lagoons and coastal salterns with NaCl concentrations between 1 and 2.6 M [53, 54]. The term “halobacteria” refers to the red-pigmented extremely halophilic Archaea, members of the family Halobacteriaceae, and the only family in the order Halobacteriales (15). Most halobacteria require 1.5 M NaCl to grow and retain the structural integrity of the cell. Halobacteria can be distinguished from halophilic bacteria by their archaeal characteristic, in particular the presence of ether-linked lipids [55]. Most halobacteria are colored red or orange due to the presence of carotenoids, but some species are colourless. Halobacteria are the most halophilic organisms known so far and form the dominant microbial population when hypersaline waters approach saturation [56, 57] (Table 1).

3 Cellulases Due to the harsh living conditions extremophiles are interesting source of stable biocatalysts. Thermostable cellulases active towards crystalline cellulose are of great biotechnological interest. Cellulose is the most abundant organic biopolymer in nature since it is the structural polysaccharide of the cell wall in the plant kingdom. It consists of glucose units linked by β-1,4-glycosidic bonds with a polymerization grade of up to 15 000 glucose units in a linear mode. The minimal molecular weight of cellulose from different sources has been estimated to vary from about 50 000 to 2 500 000 in different species, which is equivalent to 300 to 15 000 glucose residues. Although cellulose has a high affinity to water, it is completely insoluble in it. Natural cellulose compounds are structurally heterogeneous and have both amorphous and highly ordered crystalline regions. The degree of crystallinity depends

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on the source of the cellulose and the higher crystalline regions are more resistant to enzymatic hydrolysis. Cellulose can be hydrolyzed into glucose by the synergistic action of at least three different enzymes: endoglucanase (cellulase), exoglucanase (cellobiohydrolase) and β-glucosidase (cellobiase). Endoglucanase (E.C. 3.2.1.4) hydrolyzes cellulose in a random manner as endo-hydrolase producing various oligosaccharides, cellobiose and glucose. Exoglucanases (EC 3.2.1.91) hydrolyze β-1,4 D-glycosidic linkages in cellulose and cellotetraose, releasing cellobiose from the non-reducing end of the chain. β-Glucosidases (EC 3.2.1.21) catalyze the hydrolysis of terminal, nonreducing β-D-glucose residues releasing β-D-glucose. Several cellulose-degrading enzymes from various thermophilic organisms have been investigated (Table 2). A thermostable cellulase from Thermotoga maritima MSB8 has been characterized [58]. The enzyme is rather small, with a molecular weight (MW) of 27 kDa, and it is optimally active at 95 ◦ C and between pH 6.0 and 7.0 [59]. Two thermostable cellulases, CelA and CelB, with optimal activity between 95 ◦ C and 106 ◦ C, were purified from Thermotoga neapolitana [60]. Cellulase and hemicellulase genes have been found clustered together on the genome of the thermophilic anaerobic bacterium Caldocellum saccharolyticum, which grows on cellulose and hemicellulose as sole carbon sources. The gene for one of the cellulases (CelA) was isolated and was found to consist of 1751 amino acids. This is the largest cellulase gene described to date [61–63]. A large cellulolytic enzyme (CelA) with the ability to hydrolyze microcrystalline cellulose was isolated from the extremely thermophilic bacterium Anaerocellum thermophilum [61–63]. The enzyme has an apparent molecular mass of 230 kDa and exhibits significant activity towards Avicel and is most active towards soluble substrates such

Table 2 Bacterial and archaeal cellulolytic enzymes Organism

Endoglucanase

Cellobiohydrolase

β-Glucosidase

+ + + + + + + +

+ – + + – + + +

– – – – + + – –

– –

+ –

+ +

Bacteria Cellulomonas fimi Clostridium thermocellum C. stercorarium Cytophaga sp. Fibrobacter succinogenes Ruminococcus albus Thermotoga maritima Thermotoga neapolitana Archaea Pyroccus furiosus Sulfolobus solfataricus

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as carboxy-methyl-cellulose (CMC) and β-glucan. Maximal activity was observed at pH 5–6 and 85–95 ◦ C. A thermostable β-glucosidase is produced by Thermotoga sp. FjSS3-B1 [64]. The enzyme is highly thermostable and shows maximal activity at 115 ◦ C at pH 6.8–7.8. The thermostability of the enzyme is salt dependent. This enzyme is active on amorphous cellulose and carboxymethyl-cellulose. The thermophilic bacterium Rhodothermus marinus produces a hyperthermostable cellulase, with a temperature optimum of more A resolution. This is than 90 ◦ C [65], the structure of which was solved to 1.8 ˚ the first structure of a thermophilic member of family glycoside hydrolase 12 to have been solved. The beta-jelly roll fold observed has identical topology to those of the two mesophilic members of the family whose structures have been elucidated previously. A Hepes buffer molecule bound in the active site may have triggered a conformational change to an active configuration, as the two catalytic residues Glu124 and Glu207, together with dependent residues, are observed in a conformation similar to that seen in the structure of Streptomyces lividans CelB2 complexed with an inhibitor. The structural similarity between this cellulase and the mesophilic enzymes serves to highlight features that may be responsible for its thermostability, chiefly an increase in ion-pair number and the considerable stabilization of a mobile region seen in S. lividans CelB2. Additional aromatic residues in the active site region may also contribute to the difference in thermophilicity [66]. Recently, a thermostable endoglucanase, which is capable of degrading β-1,4 bonds of β-glucans and cellulose, has been identified in the archaeon Pyrococcus furiosus. The gene encoding this enzyme has been cloned and sequenced in E. coli. The purified recombinant endoglucanase hydrolyzes β-1,4 but not β-1,3 glycosidic linkages and has the highest specific activity with cellopentaose and cellohexaose as substrates [67]. In contrast to this, several β-glucosidases have been detected in archaea. In fact, archaeal β-glucosidases have been found in Sulfolobus solfataricus MT4 [68], S. acidocaldarius, S. shibatae and P. furiosus [69–71]. The enzyme from the latter microorganism is very stable and shows optimal activity at 103 ◦ C. The β-glucosidase from S. solfataricus MT4 is very resistant to various denaturants with activity up to 85 ◦ C. The gene for this β-glucosidase has been cloned and overexpressed in E. coli [72]. The less-thermoactive cellulases that are widespread in fungi and bacteria, have already found various biotechnological applications. The most effective enzyme of commercial interest is the cellulase produced by Trichoderma sp. Cellulases were also obtained from strains of Aspergillus, Penicillium and Basidomycetes [73, 74]. Cellulolytic enzymes can be used in alcohol production to improve juice yields and effective color extraction of juices. The presence of cellulases in detergents causes color brightening, softening and improves particulate soil removal [75]. Cellulase (Denimax® Novozymes) is also used for the “biostoning” of jeans instead of using stones. Other suitable applications of cellulases include the pretreatment of cellulosic biomass and forage crops to improve nutritional quality and digestibility, en-

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zymatic saccharification of agricultural and industrial wastes and production of fine chemicals [76].

4 Xylan-degrading enzymes To date only a few extreme thermophilic microorganisms are able to grow on xylan and secrete thermoactive xylanolytic enzymes (Table 3). Xylan is a heterogeneous molecule that constitutes the main polymeric compound of hemicellulose, a fraction of the plant cell wall, which is a major reservoir of fixed carbon in nature. The main chain of the heteropolymer is composed of xylose residues linked by β-1,4-glycosidic bonds. Approximately half of the xylose residues have substitution at the O-2 or O-3 positions with acetyl, arabinosyl and glucuronosyl groups. The complete degradation of xylan requires the action of several enzymes. The endoβ-1,4-xylanase (E.C.3.2.1.8) hydrolyzes β-1,4-xylosydic linkages in xylans, while β-1,4-xylosidase (EC 3.2.1.37) hydrolyzes β-1,4-xylans and xylobiose by removing the successive xylose residues from the non-reducing termini [77]. Members of the order Thermotogales and Dictyoglomus thermophilum Rt46B.1 have been described to produce xylanases that are active and stable at high temperatures [78, 79]. The most thermostable endoxylanases that have been described so far are those derived from Thermotoga sp. strain FjSS3-B.1, Thermotoga maritima [59, 80, 81], T. neapolitana [82, 83] and T. thermarum [84]). These enzymes, which are active between 80 and 105 ◦ C, are mainly cell-associated and most probably localized within the toga, which covers the cells. Several genes encoding xylanases have already been cloned and sequenced. The gene from T. maritima, encoding a thermostable xylanase has been cloned and expressed in E. coli. Comparison between the T. maritima recombinant xylanase and the commercially available enzyme, Pulpenzyme™ indicates that the thermostable xylanase could be of interest for application in pulp and paper industry [85]. Recently, Bacillus thermantarcticus, a thermophilic bacterium isolated from Antarctic geothermal soil near the crater of Mount Melbourne, was found to produce an extracellular xylanase and β-xylosidase. The optimum temperatures are 80 ◦ C for xylanase at pH 5.6 and 70 ◦ C for β-xylosidase at pH 6.0. The isoelectric points and molecular masses are 4.8 and 45 kDa for xylanase and 4.2 and 150 kDa for β-xylosidase, respectively. Xylanase is stable at 60 ◦ C for 24 h, whereas it shows a half-life at 70 ◦ C of 24 h and at 80 ◦ C of 50 min. β-Xylosidase activity does not decrease after 1 h at 60 ◦ C. Interestingly, the action of two enzymes on xylan leads to the formation of xylose [86]. The extracellular thermostable endo-1,4-β-xylanase (XT6) produced by the thermophilic bacterium Geobacillus stearothermophilus T-6 was shown to bleach pulp optimally at pH 9 and 65 ◦ C and was successfully used in a large-

ND: Not determined

Thermococcus zilligii

Archaea

Bacillus subtilis Streptomyces olivochromogenes Thermoactinomyces vulgaris Thermoanaerobacter saccharolyticum Thermonospora fusca Thermotoga maritima Thermotoga neapolitana

Bacteria

Organism

+

+ + + + + + + ND

+ + ND + + ND ND

Endoxylanase β-Xylosidase

Table 3 Bacterial and archaeal xylanolytic enzymes

ND

+ + + + + ND ND

β-L-Arabinofuranosidase

ND

+ ND ND ND ND ND ND

β-Glucuronidase

ND

+ + + + + ND ND

Acetyl Xylan esterase

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scale biobleaching mill trial. The xylanase gene was cloned and sequenced. The mature enzyme consists of 379 amino acids, with a calculated molecular weight of 43.8 kDa and a pI of 9.0. In order to study the mechanism of catalysis and to provide a structural basis for the rational introduction of enhanced thermostability by site-specific mutagenesis, the structure of wild type was refined at 2.4 ˚ A resolution. The structure demonstrates that XT6 is made up of an eightfold TIM barrel containing a deep active-site groove, consistent with its “endo” mode of action. The two essential catalytic carboxylic residues (Glu159 and Glu265) are located at the active site within 5.5 ˚ A of each other, as expected for “retaining” glycoside hydrolases. A unique subdomain was identified in the carboxy-terminal part of the enzyme and was suggested to have a role in xylan binding. The three-dimensional structure of XT6 is of great interest since it provides a favourable starting point for the rational improvement of its already high thermal and pH stabilities, which are required for a number of biotechnological and industrial applications [87]. Among the thermophilic Archaea, xylanase production has been demonstrated only in the hyperthermophilic archaeon Pyrodictium abyssi [88]. The enzyme has an optimum temperature of 110 ◦ C, which is one of the highest reported for a xylanase. Recently, an endo-1,4-xylanase and a β-xylosidase have been characterized from the extremely halophilic archaeon, Halorhabdus utahensis [89]. This is the first report on hemicellulose-degrading enzymes produced by an extremely halophilic archaeon. Xylanases from bacteria and eukarya have a wide range of potential biotechnological applications. They are already produced on industrial scale and are used as food additives in poultry, for increasing feed efficiency diets and in wheat flour for improving dough handling and the quality of baked products. In recent years, the major interest in thermostable xylanases is found in enzyme-aided bleaching of paper. The chlorinated lignin derivatives generated by this process constitute a major environmental problem caused by the pulp and paper industry. Recent investigations have demonstrated the feasibility of enzymatic treatments as alternatives to chlorine bleaching for the removal of residual lignin from pulp. Treatment of craft pulp with xylanase leads to a release of xylan and residual lignin without undue loss of other pulp components. Xylanase treatment at elevated temperatures opens up the cell wall structure, thereby facilitating lignin removal in subsequent bleaching stages [90, 91].

5 Pectin-degrading enzymes Pectin is a branched heteropolysaccharide consisting of a main chain of α-1,4-D-polygalacturonate, which is partially methyl-esterified. Along the chain, L-rhamnopyranose residues are present that are the binding sites for

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side chains composed of neutral sugars. Pectin is an important plant material that is present in the middle lamellae as well as in the primary cell walls. Pectin is degraded by pectinolytic enzymes that can be classified into two major groups. The first group comprises methylesterases, whose function is to remove the methoxy groups from pectin. The second group comprises the depolymerases (hydrolases and lyases), that attack both pectin and pectate (polygalacturonic acid). A great variety of pectinolytic bacteria have been isolated from various habitats such as trees, lakes, soil, tumen, mullet gut, and human intestinal track. Pectin hydrolases are predominantly synthesized by fungi whereas pectate lyases are mostly produced by bacteria and usually act at alkaline pH and are Ca2+ -dependent. Pectin degradation by thermophilic bacteria has been reported for Thermoanaerobacter thermohydrosulfuricus, Thermoanaerobacter thermosulfurigenes, Clostridium thermocellum, Desulfurococcus amylolyticus, Clostridium thermosaccharolyticum and Bacillus stearothermophilus [92–101]. Although many microorganisms have been screened for pectinolytic activity, little attention has been paid to pectinolytic enzymes from thermophilic and hyperthermophilic microorganisms. Previously a novel anaerobic strain from a thermal spa in Italy was isolated that produces two thermoactive lyases that have a very high affinity for polygalacturonate. This is a spore-forming anaerobic microorganism able to grow on citrus pectin and pectate optimally at 70 ◦ C, which has been identified as Thermoanaerobacter italicus. After growth on citrus pectin, two pectate lyases were induced, purified and biochemically characterized [102]. Both enzymes display similar catalytic properties and can function at temperatures up to 80 ◦ C. An increase in the enzymatic activity of both pectate lyases was observed after the addition of Ca+2 . The ability of the hyperthermophilic bacterium Thermotoga maritima to grow on pectin as a sole carbon source coincides with the secretion of a pectate lyase A (PelA) in the extracellular medium. The pel A gene of T. maritima was functionally expressed in E. coli as the first heterologously produced thermophilic pectinase, and purified to homogeneity [103]. Gel filtration indicated that the native form of PelA is tetrameric. Highest activity (422 U/mg, with a Km of 0.06 mM), was demonstrated on polygalacturonic acid (PGA), whereas pectins with an increasing degree of methylation were degraded at a decreasing rate. Similar to pectate lyases, PelA demonstrated full dependency on Ca2+ for stability and activity. The enzyme is highly thermoactive and thermostable, operating optimally at 90 ◦ C and pH 9.0, with a half-life for thermal inactivation of almost 2 h at 95 ◦ C, and an apparent melting temperature of 102.5 ◦ C. Detailed characterization of the product formation with polygalacturonic acid indicated that PelA has a unique eliminative exo-cleavage pattern liberating unsaturated trigalacturonate as the major product, in contrast with unsaturated digalacturonate for other exopectate lyases known. To date pectin-hydrolyzing enzymes from archaea have not been identified and characterized. Enzymatic pectin degradation is widely applied in food-

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technology processes, as in fruit-juice extraction, to increase the juice yield, to reduce its viscosity, improve color extraction from the skin and to macerate fruit and vegetable tissues [103].

6 Chitin-degrading enzymes Chitin is a linear β-1,4 homopolymer of N-acetyl-glucosamine residues and it is one of the most abundant natural biopolymers on earth. Particularly in the marine environment, chitin is produced in enormous amounts and its turnover is due to the action of chitinolytic enzymes. Chitin is the major structural component of most fungi and some invertebrates (crustacea and insects), while for soil or marine bacteria chitin serves as a nutrient (41). Chitin degradation is known to proceed with the endo-acting chitin hydrolase (chitinase A; EC 3.2.1.14) and the chitin-oligomer-degrading exo-acting hydrolases (chitinase B) and N-acetyl-D-glycosaminidase (trivial name: chitobiase; EC 3.2.1.52). Chitin exhibits interesting properties that make it a valuable raw material for several applications. It has been estimated that the annual worldwide formation rate and steady-state amount of chitin is of the order of 1010 to 1011 tons per year. Therefore, application of thermostable chitin-hydrolyzing enzymes (chitinases) is expected for effective utilization of this abundant biomass [104–106]. Although a large number of chitinhydrolyzing enzymes has been isolated and their corresponding genes have been cloned and characterized, only a few thermostable chitin hydrolyzing enzymes are known. These enzymes have been isolated from the thermophilic bacterium Bacillus licheniformis X-7u [107], Bacillus sp. BG-11 [108] and Streptomyces thermoviolaceus OPC-520 [109]. So far, only three hyperthermophilic archaea, Thermococcus chitonophagus [110], Thermococcus kodakaraensis KOD1 [111, 112], and Pyrococcus furiosus [113] have been shown to grow on chitin. The extreme thermophilic anaerobic archaeon Thermococcus chitonophagus has been reported to posses an enzymatic system able to hydrolyze chitin. From this microorganism, a chitinase (1,4-beta-D-N-acetylglucosaminidase, EC 3.2.1.14) was detected and purified to homogeneity in its native form. This is the first nonrecombinant chitinase purified and characterized from archaea and also constitutes the first case of a membraneassociated chitinase isolated from archaea. The enzyme is a monomer with an apparent molecular weight of 70 kDa and appears to be associated with the outer side of the cell membrane. The enzyme is optimally active at 70 ◦ C and pH 7.0 and exhibits remarkable thermostability, maintaining 50% activity even after 1 h at 120 ◦ C. The enzyme was not inhibited by allosamidin, the natural inhibitor of chitinolytic activity, and was also resistant to denaturation by urea and SDS. Chi70 shows broad substrate specificity for several chitinous substrates and derivatives and has been classified as an endochiti-

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nase due to its ability to release chitobiose from colloidal chitin [110]. Very recently, the gene encoding a chitinase from a hyperthermophilic archaeon Thermococcus kodakaraensis KOD1 was cloned, sequenced and expressed in E. coli. The purified recombinant protein is optimally active at 85 ◦ C and pH 5.0. This multidomain protein consists of two active sites with different cleavage specificities and three substrate-binding domains, which are related to two families of cellulose-binding domains. The enzyme produces chitobiose as the major end product. This thermostable chitinase, which is active in the presence of detergents and organic solvents, can be applied as useful catalyst in the industry e.g. production of N-acetyl-chitooligosaccharides with biological activity [111]. Pyrococcus furiosus was also found to grow on chitin, adding this polysaccharide to the inventory of carbohydrates utilized by this hyperthermophilic archaeon. Accordingly, two open reading frames (chiA and chiB) were identified in the genome of P. furiosus, which encode chitinases with sequence similarity to proteins from the glycosyl hydrolase family 18 in less-thermophilic organisms. The two chitinases share little sequence homology with each other, except in the catalytic region, where both have the catalytic glutamic acid residue that is conserved in all family 18 bacterial chitinases. The genes encoding ChiA, without its signal peptide, and ChiB were cloned and expressed in E. coli. The pH optima of both enzymes is about 6.0 with a broad temperature optima between 90 and 95 ◦ C. ChiA melted at 101 ◦ C, whereas ChiB was found to be extremely thermostable, with a melting temperature of 114 ◦ C. ChiA exhibited no detectable activity toward chitooligomers smaller than chitotetraose, indicating that the enzyme is an endochitinase whereas ChiB is a chitobiosidase, progressively cleaving off chitobiose from the non-reducing end of chitin or other chitooligomers. Synergistic activity was observed for the two chitinases on colloidal chitin, indicating that these two enzymes work together to recruit chitin-based substrates for P. furiosus growth. This was supported by the observed growth on chitin as the sole carbon source in a sulfur-free media [113].

7 Starch-processing enzymes 7.1 Heat-stable amylases, glucoamylases and α-glucosidases Extremely thermostable α-amylases have been characterized from a number of hyperthermophilic archaea belonging to the genera Pyrococcus [114–118] and Thermococcus [119, 120]. α-Amylase (α-1,4-glucan-4-glucanohydrolase; EC 3.2.1.1), hydrolyzes linkages in the interior of the starch polymer in a random fashion that leads to the formation of linear and branched oligosaccharides. The sugar-reducing groups are liberated in the α-anomeric config-

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uration. Most starch-hydrolyzing enzymes belong to the α-amylase family, which contains a characteristic catalytic (β α)8 -barrel domain. Throughout the α-amylase family, only eight amino acid residues are invariant, seven at the active site and a glycine in a short turn. The optimal temperatures for the activity of these enzymes range between 80 ◦ C and 100 ◦ C. Thermoactive amylolytic enzymes have also been detected in hyperthermophilic archaea of the genera Sulfolobus, Thermophilum, Desulfurococcus, and Staphylothermus [121–123] (Table 4). Molecular cloning of the corresponding genes and their expression in heterologous hosts circumvent the problem of insufficient expression in the natural host. The gene encoding an extracellular α-amylase from Pyrococcus furiosus has recently been cloned and the recomTable 4 Starch-hydrolyzing enzymes from extreme thermophilic and hyperthermophilic Archaea Enzyme

Organism (growth temperature)

Enzyme properties Optimal Optimal temperature pH

α-Amylase

Desulfurococcus mucosus (85) Pyrococcus furiosus (100) Pyrococcus sp. KOD1 (100) Pyrococcus woesei (100) Pyrodictium abyssi (98) Staphylothermus marinus (90) Sulfolobus solfatatricus (88) Thermococcus celer (85) Thermococcus profundus DT5432 (80) Thermococcus profundus (80) Thermococcus aggregans (85) Dictyoglomus thermophilum Rt46B.1 (73)

85 100 90 100 100 100

5.5 6.5–7.5 6.5 5.5 5.0 5.0

90 80 80 95 90

5.5 5.5 4.0–5.0 6.5 5.5

Pullulanase type II

Desulfurococcus mucosus (88) Pyrococcus woesei (100) Pyrodictium abyssi (98) Thermococcus celer (85) Thermococcus litoralis (90) Thermococcus hydrothermalis (80)

100 100 100 90 98 95

5.0 6.0 9.0 5.5 5.5 5.5

Pullulan-hydrolase type III

Thermococcus aggregans (85)

100

6.5

Glucoamylase

CGTase

Thermoplasma acidophilum (60) Picrophilus oshimae (60) Picrophilus torridus (60) Thermococcus sp.B1001 (75)

90 90 90 100

6.5 2.0 2.0 2.0

α-Glucosidase

Thermococcus strain AN1 (80)

130

63

Thermococcus hydrothermalis (80)

–

–

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binant enzyme has been expressed in Bacillus subtilis and E. coli (48, 49). The high thermostability of the pyrococcal extracellular α-amylase (thermal activity even at 130 ◦ C) makes this enzyme an interesting candidate for industrial application. α-Amylases with lower thermostability have been isolated from the archaea Thermococcus profundus [124], Pyrococcus kodakaraensis [125] and the bacteria Thermotoga maritima [126] and Dictioglomus thermophilum [127, 128]. The genes encoding these enzymes were successfully expressed in E. coli. Similar to the amylase from Bacillus licheniformis, which is commonly used in liquefaction of starch in the industry, the enzyme from T. maritima requires Ca2+ for activity. Further investigations have shown that the extreme marine hyperthermophilic archaeon Pyrodictium abyssi can grow on various polysaccharides and also secretes a heat-stable α-amylase [88]. Unlike α-amylase, the production of glucoamylase seems to be very rare in extremely thermophilic and hyperthermophilic bacteria and archaea (Table 4). Glucoamylases (EC 3.2.1.3) hydrolyze terminal α-1,4-linked-Dglucose residues successively from non-reducing ends of the chains, releasing β-D-glucose. Among the thermophilic anaerobic bacteria, glucoamylases have been purified and characterized from Clostridium thermohydrosulfuricum 39, Clostridium thermosaccharolyticum and Thermoanaerobacterium thermosaccharolyticum DSM 571 [129–141]. Recently, it has been shown that the thermoacidophilic archaea Thermoplasma acidophilum, Picrophilus torridus and Picrophilus oshimae produce heat- and acid-stable glucoamylases. The purified archaeal glucoamylases are optimally active at pH 2 and 90 ◦ C. Catalytic activity is still detectable at pH 0.5 and 100 ◦ C. These enzymes are more thermostable than the aforementioned glucoamylases from bacteria, yeast and fungi. This has been the first report on the production of glucoamylase in archaea [142]. However, the lack of suitable genetic methods for thermoacidophiles have precluded structural studies aimed at discovering their adaptation at very low pH. Very recently, the gene (ssg) encoding a putative glucoamylase from Sulfolobus solfataricus, was cloned and expressed in E. coli, and the properties of the recombinant protein were examined in relation to the glucose production process. This represents the first successful cloning of a glucoamylase gene from a thermoacidophilic archaeon in a mesophilic host [143]. The recombinant glucoamylase is extremely thermostable, with an optimal temperature at 90 ◦ C, however the intracellular enzyme is most active in the slightly acidic pH range from 5.5 to 6.0. The enzyme liberated β-D-glucose from the substrate maltotriose, and the substrate preference for maltotriose distinguishes this enzyme from fungal glucoamylases. Gel permeation chromatography and sedimentation equilibrium analytical ultracentrifugation analysis revealed that the enzyme exists as a tetramer [143]. The glucoamylase from S. solfataricus has a potential for improving industrial starch processing by eliminating the need to adjust both pH and temperature. However it is remarkably less

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acidic than the glucoamylases from P. torridus, P. oshima and T. acidophylum. In addition to the glucoamylase, S. solfataricus produces an α-amylase, which is secreted into the culture supernatant during growth on starch as the sole carbon and energy source. The purified enzyme is a homodimer with a subunit size of 120 kDa and catalyzes the hydrolysis of starch, dextrin, and α-cyclodextrin [123]. Assimilation of starch-derived carbon in this organism is coupled to production of a cell-associated α-glucosidase, which converts maltodextrins into glucose. S. solfataricus employs a catabolite repression (CR) system to regulate production of glycosyl hydrolases, including synthesis of the secreted α-amylase. α-Glucosidases (E C 3.2.1.20) attack the α-1,4 linkages of oligosaccharides that are produced by the action of other amylolytic enzymes. Unlike glucoamylase, α-glucosidase prefers smaller oligosaccharides, e.g. maltose and maltotriose, and liberates glucose with an α-anomeric configuration. α-Glucosidases are present in thermophilic archaea and bacteria. An intracellular and an extracellular α-glucosidase have been purified from P. furiosus and Thermococcus strain AN1 [144–148]. The enzyme exhibits optimal activity at pH 5.0–6.0 over a temperature range of 105 to 115 ◦ C; the half-life at 98 ◦ C is 48 h. An α-glucosidase (maltase) and flanking sequences from Sulfolobus solfataricus were cloned and characterized. malA is 2,083 bp and encodes a protein of 693 amino acids with a calculated mass of 80.5 kDa. It is flanked on the 5 side by an unusual 1-kb intergenic region. The purified recombinant enzyme hydrolyzes p-nitrophenyl–D-glucopyranoside with a Km of 2.16 mM and a Vmax of 3.08 µmol of p-nitrophenol/min at 85 ◦ C. It exhibited a pH optimum for maltose hydrolysis of 4.5. In contrast to its apparent greater tendency to dissociate during SDS-PAGE, the recombinant α-glucosidase exhibits greater thermostability than the native enzyme, with a half-life of 39 h at 85 ◦ C at a pH of 6.0. Unlike maltose hydrolysis, glycogen hydrolysis is optimal at the intracellular pH of the organism. These results indicate a unique role for the S. solfataricus α-glucosidase in carbohydrate metabolism [149]. 7.2 Thermoactive pullulanase and CGTase Enzymes capable of hydrolyzing α-1,6 glycosidic bonds in pullulan are defined as pullulanases. Pullulan is a linear α-glucan consisting of maltotriose units joined by α-1,6 glycosidic linkages and it is produced by Aureobasidium pullulans with a chain length of 480 maltotriose units [150]. On the basis of substrate specificity and product formation, pullulanases have been classified into two groups: pullulanase type I and pullulanase type II. Pullulanase type I (EC 3.2.1.41) specifically hydrolyzes the α-1,6-linkages in pullulan as well as in branched oligosaccharides (debranching enzyme), and its degrada-

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tion products are maltotriose and linear oligosaccharides, respectively. Pullulanase type I is unable to attack α-1,4-linkages in α-glucans. Pullulanase type II (amylopullulanase) attacks α-1,6-glycosidic linkages in pullulan and α-1,4-linkages in branched and linear oligosaccharides. The enzyme has multiple specificity and is able to fully convert polysaccharides (e.g. amylopectin) to small sugars (e.g. glucose, maltose, maltotriose) in the absence of other enzymes, such as α-amylase or β-amylase [151, 152]. Thermostable and thermoactive pullulanases from extremophilic microorganisms have been detected in Pyrococcus furiosus [153], Thermococcus celer [154], Desulfurococcus mucosus [155], Staphylothermus marinus, Thermococcus hydrothermalis [156] and Thermococcus aggregans [157] (Table 4). Temperature optima between 90 ◦ C and 105 ◦ C, as well as remarkable thermostability even in the absence of substrate and calcium ions, have been observed. Most thermoactive pullulanases identified to date belong to the type II group. Pullulanases type II from P. furiosus and P. woesei have been expressed in E. coli [153, 158, 159]. The unfolding and refolding of the pullulanase from P. woesei has been investigated using guanidinium chloride as denaturant. The monomeric enzyme (90 kDa) was found to be very resistant to chemical denaturation and the transition midpoint for guanidinium chloride-induced unfolding was determined to be 4.8. The unfolding process was reversible. Reactivation of the completely denatured enzyme (in 7.8 M guanidinium chloride) was obtained upon removal of the denaturant by stepwise dilution, 100% reactivation was observed when refolding was carried out via a guanidinium chloride concentration of 4 M in the first dilution step [160]. On the basis of the amino-acid sequence, the pullulanase type II (amylopullulanase) from T. hydrothermalis and P. furiosus belong to family 57 (GH-57) of the glycoside hydrolases. Five conserved regions were identified, which are postulated to be GH-57 consensus motifs by comparison to the 659amino-acid-long 4–glucanotransferase from Thermococcus litoralis. These motifs correspond to 13_HQP (region I), 76_GQLEIV (region II), 120_WLTERV (region III), 212_HDDGEKFGVW (region IV), and 350_AQCNDAYWH (region V). The third and fourth conserved regions contain the hypothetical catalytic nucleophile E291 and the proton donor D394, respectively. To validate this prediction, the characterization of catalytic sites of certain members of GH-57 has been started recently. Site-directed mutagenesis performed by Zona et al. [161] on pullulanase type II from T. hydrothermalis reveals that both residues are indeed critical for the pullulanolytic and amylolytic activities of the pullulanase type II [161]. The crucial role of E291 as the catalytic nucleophile has also been confirmed by Kang et al. [162], who performed similar experiment on the pullulanase type II from P. furiosus (PfAPU). The apparent catalytic efficiencies (kcat /Km ) of mutants E291Q and D394N on pullulan were 123.0 and 24.4 times lower, respectively, than that of PfAPU. The activity of mutant E396Q on pullulan was too low to allow reliable determination of its catalytic efficiency. The apparent specific activities of these

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enzymes on starch also decreased 91.0 times (E291Q), 11.7 times (D394N), and 37.2 times (E396Q). The hydrolytic patterns for pullulan and starch were the same, while the hydrolysis rates differed as reported. Therefore, these data support the prediction and strongly suggest that the biofunctionality of the pullulanase type II is determined by a single catalytic centre. Interestingly, pullulanase type I has not been isolated in Archaea so far, whereas the enzyme has been characterized in several thermophilic microorganisms. The aerobic thermophilic bacterium Thermus caldophilus GK-24 produces a thermostable pullulanase of type I when grown on starch [163]. The pullulanase is optimally active at 75 ◦ C and pH 5.5, is thermostable up to 90 ◦ C, and does not require Ca2+ for either activity or stability. The first debranching enzyme (pullulanase type I) from an anaerobic thermophile was identified in the bacterium Fervidobacterium pennivorans Ven5 which was cloned and expressed in E. coli. The enzyme from F. pennivorans Ven5 attacks exclusively the α-1,6-glycosidic linkages in polysaccharides [164, 165]. This thermostable debranching enzyme leads to formation of long-chain linear polysaccharides from amylopectin [166]. The same enzyme has been also characterized from the related microorganism T. maritima [167]. Interestingly, data concerning the physico-chemical properties of all debranching enzymes reported so far show that they are mostly active in the acidic or neutral pH range. Until very recently, no reports have been present on the ability of thermophilic microorganisms to produce heat and alkaline stable pullulanase type I. After sequencing the whole genome of the thermoalkaliphile A. gottschalkii, an open reading frame with high pairwise similarity to the pullulanases from the thermophilic anaerobic bacteria F. pennivorans and T. maritima was identified and the gene (encoding 865 amino acids with a predicted molecular mass of 98 kDa) was cloned and expressed in E. coli. Pullulan hydrolysis activity was optimal at pH 8.0 and 70 ◦ C, and under these physicochemical conditions the half-life of rPulAg was 22 h. The pullulanase from A. gottschalkii, therefore, is the first thermoalkalistable type I pullulanase that has been described [168]. Thermostable cyclodextrin glycosyltransferases (CGTases) are produced by Thermoanaerobacter species [169], Thermoanaerobacterium thermosulfurigenes [170] and Anaerobranca gottschalkii (13, 66, 67). Cyclodextrin glycosyltransferase (CGTase, EC 2.4.1.19) is an enzyme that is generally found in bacteria and was recently discovered in archaea. The archaeal enzyme was found in Thermococcus sp. and is optimally active at 100 ◦ C (Table 4). This enzyme produces a series of non-reducing cyclic dextrins from starch, amylose, and other polysaccharides. α-, β- and γ -cyclodextrins are rings formed by 6, 7, and 8 glucose units, respectively, that are linked by α-1,4-bonds [171]. The finding of extremely thermophilic bacteria and archaea capable of producing novel thermostable starch-hydrolyzing enzymes is a valuable contribution to the starch-processing industry. By using robust starch-modifying enzymes from thermophiles, innovative and environmentally friendly pro-

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cesses can be developed, aiming at the formation of products of high added value for the food industry. At elevated temperatures starch is more soluble (30 to 35% w/v) and the risk of contamination is reduced. This is of advantage when starch will be converted to high-glucose and high-fructose syrups. Industrial production of fructose from starch consists of three steps: liquefaction, saccharification and isomerization. This multistage process (step 1: pH 6.5, 98 ◦ C; step 2: pH 4.5, 60 ◦ C: step 3; pH 8.0, 65 ◦ C) leads to the conversion of starch to fructose with concurrent formation of high concentrations of salts that have to be removed by ion exchangers. Furthermore, high energy is required for cooling from 100 ◦ C to 60 ◦ C in step 2. The application of thermostable enzymes such as amylases, glucoamylases, pullulanases and glucose isomerases that are active and stable above 100 ◦ C and at acidic pH values can simplify this complicated process. Therefore, strong efforts have been invested in the isolation of thermostable and thermoactive amylolytic enzymes from hyperthermophiles, since they could improve the starch conversion process and lower the cost of sugar-syrup production. The use of the extremely thermostable amylolytic enzymes can lead to other valuable products, which include innovative starch-based materials with gelatine-like characteristics and defined linear dextrins that can be used as fat substitutes, texturizers, aroma stabilizers and prebiotics. CGTases are used for the production of cyclodextrins that can be used as a gelling, thickening or stabilizing agent in jelly desserts, dressing, confectionery, dairy and meat products. Due to the ability of cyclodextrins to form inclusion complexes with a variety of organic molecules, cyclodextrins improve the solubility of hydrophobic compounds in aqueous solution. This is of interest for the pharmaceutical and cosmetic industries. Cyclodextrin production is a multistage process in which starch is first liquefied by a heat-stable amylase followed by a second step in which a less-thermostable CGTase from Bacillus sp. is used. The application of heatstable CGTase in jet cooking, where temperatures up to 105 ◦ C are achieved, will allow liquefaction and cyclization to take place in one step [172].

8 Proteolytic enzymes Proteases are involved in the conversion of proteins to amino acids and peptides. They have been classified according to the nature of their catalytic site in the following groups: serine, cysteine, aspartic, or metallo proteases. A variety of heat-stable proteases has been identified in hyperthermophilic archaea belonging to the genera Desulfurococcus [173], Sulfolobus [174, 175], Staphylothermus [176], Thermococcus [177, 178], Pyrobaculum [179] and Pyrococcus [180, 181] (Table 5). It has been found that most proteases from extremophiles belong to the serine type, are stable at high temperatures even in the presence of high concentrations of detergents and denaturing agents

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Table 5 Properties of thermoactive proteolytic enzymes from extreme thermophilic and hyperthermophilic Archaea Protease

Organism (growth temperature)

Enzyme properties Optimal Optimal temperature pH

Serine protease

Desulfurococcus mucosus (85) Pyrococcus furiosus (100) Pyrobaculum aerophilum (95) Thermococcus aggregans (75) Thermococcus celer (85) Thermococcus litoralis (90) Thermococcus stetteri (75) Staphylothermus marinus (90) Sulfolobus solfataricus (88)

95 8.5 – 90 95 95 85 – –

7.5 6.3 – 7.0 7.5 9.5 8.5 9.0 6.5–8

Thiol protease

Pyrococcus sp. KOD1 (95)

110

7

Acidic protease

Sulfolobus acidocaldarius (70)

Aminopeptidase I Aminopeptidase II Endopeptidase I,II,III Carboxypeptidase

Sulfolobus solfataricus (88)

90 – – – –

2.0 – – – –

(59, 68, 69). A heat-stable serine protease was isolated from the cell-free supernatant of the hyperthermophilic archaeon Desulfurococcus strain Tok12 S1 . Recently, a cell-associated serine protease was characterized from the Desulfurococcus strain SY that has a half-life of 4.3 h at 95 ◦ C [173]. A globular serine protease from Staphylothermus marinus was found to be extremely thermostable. The properties of extracellular serine proteases from a number of Thermococcus species have been analyzed [177, 178]. The extracellular enzyme from T. stetteri has a molecular mass of 68 kDa and is highly stable and resistant to chemical denaturation, as illustrated by a half-life of 2.5 h at 100 ◦ C and retention of 70% of its activity in the presence of 1% SDS [182]. A novel intracellular serine protease (pernisine) from the aerobic hyperthermophilic archaeon Aeropyrum pernix K1 was purified and characterized. At 90 ◦ C, the enzyme has a broad pH profile and an optimum at pH 9.0 for peptide hydrolysis [183, 184]. The pernisine, lacking the leader sequence, was expressed in E. coli as a fusion protein with glutathione-Stransferase. The biochemical properties of the recombinant enzyme were found to be similar to those of the native enzyme [185]. Several proteases from hyperthermophiles have been cloned and sequenced but, in general, their expression in mesophilic hosts is difficult. A gene encoding a subtilisinlike serine protease, named aereolysin has been cloned from Pyrobaculum aerophilum and the protein was modeled based on structures of subtilisintype proteases [184]. Multiple proteolytic activities have been observed in P.

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furiosus. The cell-envelope-associated serine protease of P. furiosus called pyrolysin was found to be highly stable with a half-life of 20 min at 105 ◦ C [186]. The pyrolysin gene was cloned and sequenced and it was shown that this enzyme is a subtilisin-like serine protease [187, 188]. A serine protease from Aquifex pyrophilus was cloned and weakly expressed in E coli as active and processed forms. The activity of the enzyme was highest at 85 ◦ C and pH 9. The half-life of the protein (6 h at 105 ◦ C) makes it one of the most heat-stable protease known to date [189]. Proteases have also been characterized from the thermoacidophilic archaea Sulfolobus solfataricus [175] and S. acidocaldarius [190–192]. In addition to the serine proteases other types of enzymes have been identified in extremophiles: a thiol protease from Pyrococcus sp. KOD1, a propylpeptidase (PEPase) and a new type of protease from P. furiosus [178, 181, 193]. Thermostable serine proteases were also detected in a number of extreme thermophilic bacteria blonging to the genera Thermotoga and Fervidobacterium. The enzyme system from Fervidobacterium pennivorans is able to hydrolyze feather keratin-forming high-value products such as amino acids and peptides. The enzyme, which has been named fervidolysin, is optimally active at 80 ◦ C and pH 10.0 [194]. The gene encoding fervidolysin was isolated using degenerate primers combined with Southern hybridization and inverse polymerase chain reaction. Amino-terminal-sequence analysis of these bands and their comparison with that determined from biochemically purified keratinase and its predicted protein sequence identified them as a 73-kDa fervidolysin precursor, a 58-kDa mature fervidolysin, and a 14-kDa fervidolysin propeptide. Using site-directed mutagenesis, the active-site histidine residue at position 79 was replaced by an alanine residue. The resulting fervidolysin showed a single protein band corresponding in size to the 73-kDa fervidolysin precursor, indicating that its proteolytic cleavage resulted from an autoproteolytic process. Assays using keratin and other proteinaceous substrates did not display fervidolysin activity, perhaps because of the tight binding of the propeptide in the substrate-binding site, where it could then function as an inhibitor. Finally, the recombinant fervidolysin has been crysA resolution. The tallized and the structure has been determined at 1.7-˚ crystal structure shows that the protease is composed of four domains: a catalytic domain (CD), two beta-sandwich domains (SDs), and the PD domain. A structural alignment shows a distant relationship between the PD–CD substructure of fervidolysin and pro-subtilisin E. Tight binding of PD to the remaining part of the protease is mediated by hydrogen bonds along the domain surfaces and around the active cleft, and by the clamps to SD1 and SD2 [195, 196]. The amount of proteolytic enzymes produced worldwide on a commercial scale is the largest compared to the other biotechnological enzymes in use. Serine alkaline proteases are used as additives to household detergents for laundering, where they have to resist denaturation by detergents and al-

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kaline conditions. Proteases showing high keratinolytic activity are used for soaking in the leather industry. Proteases are also used as catalysts for peptide synthesis using their reverse reaction.

9 Lipases Lipases, triacylglycerol hydrolases, are an important group of biotechnologically relevant enzymes and they find immense applications in the food, dairy, detergent and pharmaceutical industries. Lipases are produced by microbes and specifically bacterial lipases play a vital role in commercial ventures. Some important lipase-producing bacterial genera include Bacillus, Pseudomonas and Burkholderia. Lipases are generally produced on lipidic carbon, such as oils, fatty acids, glycerol or tweens in the presence of an organic nitrogen source. Bacterial lipases are mostly extracellular and are produced by submerged fermentation. The enzyme is most commonly purified by hydrophobic interaction chromatography, in addition to some modern approaches such as reverse micellar and aqueous two-phase systems. Most lipases can act in a wide range of pH and temperature, though alkaline bacterial lipases are more common. Bacterial lipases generally have temperature optima in the range 30–60 ◦ C [197]. However, highly thermotolerant lipases have been reported from B. stearothermophilus, with a half-life of 15–25 min at 100 ◦ C [198] and B. thermoleovorans [199, 200]. Lipases are serine hydrolases and have high stability in organic solvents. Besides these, some lipases exhibit chemo-, regio- and enantioselectivity. Very recently, more than five anaerobic thermophilic bacteria were found to produce extremely heat-stable lipases. They are active at a broad temperature (50–95 ◦ C) and pH (3–11) (unpublished results, Antranikian). The latest trend in lipase research is the development of novel and improved lipases through molecular approaches such as directed evolution and exploring natural communities by the metagenomic approach. The recent determination of structure of Bacillus stearothermophilus P1 lipase provides a template for other thermostable lipases, and offers insight into mechanisms used to enhance thermal stability which may be of commercial value in engineering lipases for industrial uses [201].

10 Glucose isomerases, alcohol dehydrogenases and esterases In addition to the described extracellular enzymes, intracellular enzymes from extremophiles are of interest for various applications (Table 6). Glucose isomerase or xylose isomerase (D-xylose ketol-isomerase; EC 5.3.1.5) cat-

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Table 6 Enzymes with potential biotechnological application from extreme thermophilic and hyperthermophilic archaea and bacteria Enzyme

Organism (growth temperature)

Biocatalysis

α-Amylase

Pyrococcus woesei (100 ◦ C)

Hydrolysis of α-1,4 Starch industry, bioglycosidic linkages conversion of starch in starch to glucose syrup

Debranching enzyme (pullulanase type I)

Fervidobacterium pennivorans Ven5 (75 ◦ C)

Debranching of amylopectin to linear oligosaccharides

CGTase

Thermococcus sp. Production of (75 ◦ C), Anaerobranca cyclodextrins gottschalkii (60 ◦ C)

Gelling, thickening, stabilizing agents in food industry

Cellulase

Pyrococcus furiosus (100 ◦ C)

Color extraction of juice, color brightening, improving nutritional quality

Endoxylanases

Thermotoga maritima Degradation of xylan MSB8 (80 ◦ C)

Bleaching of paper

Chitinase

Thermococcus kodakaraensis (95 ◦ C)

Degradation of chitin

Utilization of biomass of marine environment

Serine protease

Fervidobacterium pennivorans (70 ◦ C)

Keratin hydrolysis

Soaking in leather industry, production of amino acids and peptides from feathers

Glucose isomerase

Thermotoga maritima (80 ◦ C)

Isomerization of glucose to fructose

Production of highfructose corn syrup

Alcohol deydrogenase

Sulfolobus solfataricus (88 ◦ C)

Oxidation of Reduction of ketones secondary alcohols

Esterase

Sulfolobus tokadaii (100 ◦ C)

Cleavage of esters

Biotransformation in organic solvents

DNA synthesis

Taq polymerase, polymerase chain reaction (PCR)

DNA polymerase Thermus aquaticus (75 ◦ C)

Hydrolysis of cellulose to glucose

Application

Starch bioconversion to glucose

alyzes the reversible isomerization of D-glucose and D-xylose to D-fructose and D-xylulose, respectively. The enzyme has the largest market in the food industry because of its application in the production of high-fructose corn syrup (HFCS). HFCS, an equilibrium mixture of glucose and fruc-

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tose, is 1.3 times sweeter than sucrose. Glucose isomerase is widely distributed in mesophilic microorganisms. Intensive research efforts are directed toward improving the suitability of glucose isomerase for industrial application. To reach fructose concentration of 55% the reaction must approach 110 ◦ C. Improved thermostable glucose isomerases have been engineered from mesophilic enzymes [202]. The gene encoding a xylose isomerase (XylA) of Thermus flavus AT62 was cloned and the DNA sequence was determined. XylA has an optimum temperature at 90 ◦ C and pH 7.0; divalent cations such as Mn2+ , Co2+ and Mg2+ are required for enzyme activity [203]. Thermoanaerobacterium strain JW/SL-YS 489 forms a xylose isomerase, which is optimally active at pH 6.4 and 60 ◦ C or pH 6.8 and 80 ◦ C. Like other xylose isomerases, this enzyme requires Mn2+ , Co2+ and Mg2+ for thermal stability (stable for 1 h at 82 ◦ C in the absence of substrate). The gene encoding the xylose isomerase of the Thermoanaerobacterium strain JW/SL-YS 489 was cloned and expressed in E. coli [204]. Comparison of the deduced amino acid sequence with sequences of other xylose isomerases showed that the enzyme has 98% homology with a xylose isomerase from a closely related bacterium Thermoanaerobacterium saccharolyticum B6ARI. A thermostable glucose isomerase was purified and characterized from Thermotoga maritima. This enzyme is stable up to 100 ◦ C, with a half-life of 10 min at 115 ◦ C [205]. Interestingly, the glucose isomerase from Thermotoga neapolitana displays a catalytic efficiency at 90 ◦ C, which is 2 to 14 times higher than any other thermoactive glucose isomerases at temperatures between 60 ◦ C and 90 ◦ C [205–208]. Dehydrogenases are enzymes belonging to the class of oxidoreductases. Within this class, alcohol dehydrogenases (E.C.1.1.1.1, also named ketoreductases) represent an important group of biocatalysts due to their ability to stereospecifically reduce prochiral carbonyl compounds. Alcohol dehydrogenases can be used efficiently in the synthesis of optically active alcohols, which are key building blocks for the fine-chemicals industry [209]. From a practical point of view, alcohol dehydrogenases that use nicotinamide adenine dinucleotide (NADH) as cofactor are of particular importance, because the formate/formate dehydrogenase system represents an established method to regenerate NADH efficiently. By contrast, for nicotinamide adenine dinucleotide phosphate (NADP)-dependent enzymes the cofactor-recycling systems that are available are much less efficient. The secondary specific alcohol dehydrogenase (ADH), which catalyzes the oxidation of secondary alcohols and, less readily, the reverse reaction (the reduction of ketones) has a promising future in biotechnology [209]. Although ADHs are widely distributed among microorganisms, only few examples derived from hyperthermophilic microorganisms are currently known. Among the extreme thermophilic bacteria, Thermoanaerobacter ethanolicus 39E was shown to produce an ADH, whose gene was cloned and expressed in E. coli. Interestingly, a mutant has been found to posses an advantage over the wild-type enzyme by using the

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more-stable cofactor NAD instead of NADP [210–213]. An alcohol dehydrogenase (ADH) was purified from an extremely thermophilic bacterium, Thermomicrobium roseum. The native enzyme is a homodimer of 43-kDa subunits. The pI of the enzyme was determined to be 6.2, while its optimum pH is 10.0. The enzyme oxidized mainly primary aliphatic alcohols and exhibited high substrate specificity towards ethanol, n-propanol and crotyl alcohol. The highest reaction rate was observed when ethanol was used as substrate and the Km value of the enzyme for ethanol was 24.2 mM. Pyrazole notably inhibited the enzymatic activity. The enzyme had an optimal temperature of 70 ◦ C and is highly stable [214, 215]. In addition, a novel NADH-dependent alcohol dehydrogenase (RE-ADH) was found in Rhodococcus erythropolis. This ADH catalyzes the reduction of ketones to the corresponding (S)-alcohols. The enzyme was purified and its biochemical characteristics were investigated. The gene encoding for 348 amino acids was cloned in E. coli cells and successfully expressed [216]. The subunit molecular mass as deduced from the amino acid sequence was determined to be 36 kDa. The recombinant enzyme exhibits high thermostability, which facilitated its purification by heat treatment, followed by two column-chromatography steps. RE-ADH shows high similarity to several zinc-containing medium-chain alcohol dehydrogenases. All zinc ligands seem to be conserved except for one of the catalytic zinc ligands, where Cys is probably substituted by Asp. In extreme thermophilic archaea, ADHs have been studied from Sulfolobus solfataricus and Thermococcus stetteri [217, 218]. The enzyme from S. solfataricus requires NAD as a cofactor and contains Zn ions. In contrast to ADHs from bacteria and Eukarya, the enzyme from T. stetteri lacks metal ions. The enzyme catalyzes preferentially the oxidation of primary alcohols, using NADP as cofactor and it is very thermostable, showing half-lives of 15 min at 98 ◦ C and 2 h at 85 ◦ C. Compared to mesophilic enzymes, the ADH from T. litoralis represents a new type of alcohol-oxidizing enzyme system [217, 218]. The gene of ADHs from S. solfataricus was expressed in E. coli and characterized. In the field of biotechnology, esterases are gaining increasing attention because of their application in organic biosynthesis. In aqueous solution, esterases catalyze the hydrolytic cleavage of esters to form the constituent acid and alcohol, whereas in organic solutions, the transesterification reaction is promoted. Both the reactants and the products of transesterification are usually highly soluble in the organic phase and the reactants may even form the organic phase themselves. The lipA gene encoding a thermostable esterase was cloned from Thermoanaerobacter tengcongensis and overexpressed in E. coli. The recombinant esterase, with a molecular mass of 43 kDa, hydrolyzes tributyrin but not olive oil. The esterase was optimally active at 70 ◦ C (over 15 min) and at pH 9. It is highly thermostable, with a residual activity greater than 80% after incubation at 50 ◦ C for more than 10 h [219]. An esterase from the putative esterase gene selected from the total genome analysis from the thermoacidophilic archaeon Sulfolobus tokodaii strain 7 was

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cloned and expressed as a fusion protein in E. coli. The optimum activity for ester cleavage against p-nitrophenyl esters was observed at around 70 ◦ C and pH 7.5–8.0. The enzyme exhibits high thermostability and also shows activity in a mixture of a buffer and water-miscible organic solvents, such as acetonitrile and dimethyl sulfoxide. From the kinetic analysis, p-nitrophenyl butyrate was found to be a better substrate than caproate and caprylate [220]. The Pyrococcus furiosus esterase gene was cloned in E. coli and the functional properties were determined. The archaeal enzyme is the most thermostable (a half-life of 50 min at 126 ◦ C) and thermoactive (temperature optimum 100 ◦ C) esterase known to date [221].

11 Amidases Amidases [EC 3.5.1.4] catalyze the conversion of amides to the corresponding carboxylic acids and ammonia, according to the following reaction: RCONH2 + H2 O → RCOOH + NH3 . A number of amidases from bacteria have been purified and characterized and the crystal structure of the peptide amidase from Stenotrophomonas maltophilia [222] was also resolved recently. Amidases exhibit different physicochemical characteristics and a diverse substrate spectrum. The enzymes that lack the characteristic signature GGSS(GAS)-S [223] in the primary structure are only able to hydrolyze aliphatic substrates. In contrast, amidases containing this signature are able to convert cyclic and aromatic amides as well [224–228]. These amidases are highly S-enantioselective, usually forming the optically pure acids with an enantiomeric excess above 99% [229]. Only in chemical and pharmaceutical industries for the production of optically pure compounds [230–232], drugs [225, 233], acrylic [234] and hydroxamic acids [235]. In general the application of efficient biocatalysts in industrial processes allows the formation of highly pure products with a concomitant reduction of wastes [233]. Running such processes at elevated temperatures also has many advantages, including significant improvement of transfer rates, higher substrate solubility and reduced risk of contamination. Very little, however, is known about amidases that are active at high temperatures. Three thermoactive amidases described so far were isolated from Brevibacillus borstelensis BCS-1 [236], Klebsiella pneumoniae NCTR 1 [237] and Sulfolobus solfataricus [238]. Very recently, the first thermoactive and thermostable amidase from the thermophilic actinomycete Pseudonocardia thermophila has been purified and characterized [239]. The amidase is active over a broad pH (pH 4–9) and temperature range (40–80 ◦ C) and has a half-life of 1.2 h at 70 ◦ C. The amidase has a broad substrate spectrum, including aliphatic, aromatic and amino acid amides. The presence of a double bond or a methyl group near the carboxamide group of aliphatic and amino

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acid amides enhances the enzymatic activity. The highest acyl transferase activity was detected with hexanoamide, isobutyramide and propionamide. The amidase is highly S-stereoselective for 2-phenylpropionamide; and the racemic amide was converted to the corresponding S-acid with an enantiomeric excess of > 95% at 50% conversion of the substrate. In contrast, the d,l-tryptophanamide and d,l-methioninamide were converted to the corresponding d,l-acids at the same rate.

12 DNA-processing enzymes 12.1 Polymerase chain reaction (PCR) DNA polymerases (EC 2.7.7.7) are the key enzymes in the replication of cellular information present in all life forms. They catalyze, in the presence of Mg2+ -ions, the addition of a deoxyribonucleotide 5 -triphosphate onto the growing 3 -OH end of a primer strand, forming complementary base pairs to a second strand. More than 100 DNA polymerase genes have been cloned and sequenced from various organisms, including thermophilic bacteria and archaea [256]. Thermostable DNA polymerases play a major role in a variety of molecular biological applications, e.g. DNA amplification, sequencing or labelling (Table 7). One of the most important advances in molecular biology during the last ten years is the development of polymerase chain reaction (PCR) [257–259]. The first described PCR procedure utilized the Klenow fragment of E. coli DNA polymerase I, which was heat-labile and had to be added during each cycle following the denaturation and primer-hybridization steps. Introduction of thermostable DNA polymerases in PCR facilitated the automation of the thermal cycling part of the procedure. The DNA polymerase I from the bacterium Thermus aquaticus, called Taq polymerase, was the first thermostable DNA polymerase characterized and applied in PCR. The Taq polymerase has a 5 -3 -exonuclease activity, but no detectable 3  5 -exonuclease activity. Due to the absence of a 3 -5 -exonuclease activity, this enzyme is unable to excise mismatches and, as a result, the base insertion fidelity is low. The use of high-fidelity DNA polymerases is essential for reducing the increase of amplification errors in PCR products that will be cloned, sequenced and expressed. Several thermostable DNA polymerases with 3 -5 -exonuclease-dependent proofreading activity have been described and the error rates (number of misincorporated nucleotides per base synthesized) for these enzymes have been determined. A thermostable DNA polymerase from Thermotoga maritima was reported to have a 3 -5 -exonuclease activity [260]. Archaeal proofreading polymerases such as Pwo pol

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Table 7 The most commonly used thermoactive DNA polymerases Enzyme

Organism of origin

Bacterial DNA polymerases Taq pol I Tth pol Tfi pol Tfl pol Tca pol BstI pol Tma pol

Thermus aquaticus Thermus thermophilus Thermus filiformis Thermus flavus Thermus caldophilus GK24 Bacillus stearothermophilus Thermotoga maritima

Archaeal DNA polymerases Pwo pol Pfu pol DeepVent pol KOD1 pol Vent pol 9◦ N-7 pol

Pyrococcus woesei Pyrococcus furiosus Pyrococcus sp. GB-D Pyrococcus sp. KOD1 Thermococcus litoralis Thermococcus sp. 9◦ N-7

from Pyrococcus woesei, Pfu pol from Pyrococcus furiosus, Deep Vent™ pol from Pyrococcus strain GB-D [261–268] or Vent™ pol from Thermococcus litoralis [269–274] have an error rate that is up to ten times lower than that of Taq polymerase. The 9◦ N-7 has a fivefold higher 3 -5 -exonuclease activity than T. litoralis DNA polymerase. However, Taq polymerase was not replaced by these DNA polymerases because of their low extension rates among other factors. DNA polymerases with higher fidelity are not necessarily suitable for amplification of long DNA fragments because of their potentially strong exonuclease activity. The recombinant KOD1 DNA polymerase from Thermococcus kodakaraensis KOD1 has been reported to show low error rates (similar values to those of Pfu), high processivity (persistence of sequential nucleotide polymerization) and high extension rates, resulting in an accurate amplification of target DNA sequences up to 6 kb [275–278]. To optimize the delicate competition of polymerase and exonuclease activity, the exo-motif of the The 9◦ N-7 DNA polymerase was mutated in an attempt to reduce the level of exonuclease activity without totally eliminating it. An additional problem in the performance of PCR is the generation of nonspecific templates prior to thermal cycling. Several approaches have been made to prevent the elongation of polymerase before cycling temperatures are reached. As well as using wax as a mechanical barrier between DNA and the enzyme, more sophisticated methods were invented such as the inhibition of Taq polymerase by a neutralizing antibody at mesophilic temperatures or heat-mediated activation of the immobilized enzyme.

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Recently, the PCR technique has been improved to allow low-error synthesis of long amplificates (20–40 kb) by adding small amounts of thermostable, archaeal proofreading DNA polymerases, containing 3 -5 -exonuclease activity, to Taq or other non-proofreading DNA polymerases. In this long PCR, the reaction conditions are optimized for long extension by adding different components such as gelatin, Triton X-100 or bovine serum albumin to stabilize the enzymes and mineral oil to prevent evaporation of water in the reaction mixture. In order to enhance specificity, glycerol or formamide are added. 12.2 DNA sequencing DNA sequencing by the Sanger method [279] has undergone countless refinements in the last 20 years. A major step forward was the introduction of thermostable DNA polymerases leading in the cycle sequencing procedure. This method uses repeated cycles of temperature denaturation, annealing and extension with dideoxy-termination to increase the amount of sequencing product by recycling the template DNA. Due to this “PCR-like” amplification of the sequencing products several problems could be overcome. Caused by the cycle denaturation, only fmoles of template DNA are required, no separate primer annealing step is needed and unwanted secondary structures within the template are resolved at high-temperature elongation. The first enzyme used for cycle sequencing was the thermostable DNA polymerase I from Thermus aquaticus [280, 281]. As described by Longley et al. (118) the enzyme displays 5 -3 -exonuclease activity that is undesirable because of the degradation of sequencing fragments. A combination of thermostable enzymes has been developed that produces higher quality cycle sequences. Thermo Sequenase DNA polymerase is a thermostable enzyme engineered to catalyze the incorporation of ddNTPs with an efficiency several thousand times better than other thermostable DNA polymerases. Since the enzyme also catalyzes pyrophosphorolysis at dideoxy termini, a thermostable inorganic pyrophosphatase is needed to remove the pyrophosphate produced during sequencing reactions. Thermoplasma acidophilum inorganic pyrophosphatase (TAP) is thermostable and effective for converting pyrophosphate to orthophosphate. The use of the combination of Thermo Sequenase polymerase and TAP for cycle sequencing yields sequence data with uniform band intensities, allowing the determination of longer, more accurate sequence reads. Uniform band intensities also facilitate interpretation of sequence anomalies and the presence of mixed templates. Sequencing PCR products of DNA amplified from heterozygous diploid individuals results in signals of equal intensity from each allele [282].

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12.3 Ligase chain reaction A variety of analytical methods are based on the use of thermostable ligases. Of considerable potential is the construction of sequencing primers by high-temperature ligation of hexameric primers [283], the detection of trinucleotide repeats through repeat expansion detection (RED) [284] or DNA detection by circularization of oligonucleotides [285]. Over the years several additional thermostable DNA ligases were discovered. Bacterial enzymes were derived and cloned from Thermus scotoductus and Rhodothermus marinus [286–289]. Recent studies in the crude extract of 103 strains of the genera Thermus, Bacillus, Rhodothermus and Hydrogenobacter revealed the presence of thermostable DNA ligases in 23 of the Thermus strains [290]. To date an archaeal DNA ligase has been described from Desulfurolobus ambivalens [291]. Unlike bacterial enzymes, this ligase is NAD+ -independent but ATP-dependent similar to the enzymes from bacteriophages, eukaryotes and viruses. A gene encoding DNA ligase (ligTk) from a hyperthermophilic archaeon, Thermococcus kodakaraensis KOD1, has been cloned and sequenced, and its protein product has been characterized: ligTk consists of 1686 bp, corresponding to a polypeptide of 562 amino acids with a predicted molecular mass of 64 079 Da. Sequence comparison with previously reported DNA ligases and the presence of conserved motifs suggested that ligTk was an ATP-dependent DNA ligase. Phylogenetic analysis indicated that ligTk was closely related to the ATP-dependent DNA ligase from Methanobacterium thermoautotrophicum H, a moderately thermophilic archaeon, along with putative DNA ligases from Euryarchaeota and Crenarchaeota. Recombinant ligTk was monomeric, as is the case for other DNA ligases. The protein displayed DNA ligase activity in the presence of ATP and Mg2+ . The optimum pH of ligTk was 8.0, the optimum concentration of Mg2+ , which was indispensable for the enzyme activity, was 14 to 18 mM, and the optimum concentration of K+ was 10 to 30 mM. ligTk did not display single-stranded DNA ligase activity. At enzyme concentrations of 200 nM, significant DNA ligase activity was observed even at 100 ◦ C. Unexpectedly, ligTk displayed a relatively small, but significant, DNA ligase activity when NAD+ was added as cofactor. Treatment of NAD+ with hexokinase did not affect this activity, excluding the possibility of contaminant ATP in the NAD+ solution. This unique cofactor specificity was also supported by the observation of adenylation of ligTk with NAD+ . This was the first biochemical study of a DNA ligase from a hyperthermophilic archaeon [292]. The ability of DNA ligases to use either ATP or NAD+ as a cofactor appears to be specific to DNA ligases from Thermococcales. An archaeal DNA ligase has been cloned from Thermococcus fumicolans (Tfu). The optimum temperature and pH of Tfu DNA ligase were 65 ◦ C and 7.0, respectively. The optimum concentration of MgCl2 , which is indispensable for the enzyme activity, was 2 mM. Tfu DNA ligase dis-

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plays nick joining and blunt-end ligation activity using either ATP or NAD+ , as a cofactor [293].

13 Conclusion Owing to their properties such as activity over a wide temperature and pH range, substrate specificity, stability in organic solvents, diverse substrate range and enantioselectivity, biocatalysts from extremophilic microorganisms will represent the choice for countless future applications in industry. Their importance is increasing daily in several fields, such as food additives, detergent industry, chemicals, pharmaceuticals, etc. The growing demand for more robust biocatalysts has shifted the trend towards improving the properties of existing proteins for established industrial processes and producing new enzymes that are tailor-made for entirely new areas of application. Currently, the number of novel microbial extremophilic enzymes being cloned and biochemically characterized is steadily increasing. New technologies such as genomics, metanogenomics, gene shuffling, and DNA evolution provide valuable tools for improving or adapting enzyme properties to the desired requirements. However, the success of these techniques demands the production of recombinant enzymes at a high level, allowing experimental trials and application tests. Thus, the modern methods of genetic engineering combined with an increasing knowledge of structure, function relationship and process engineering will allow further adaptation of enzymes to industrial needs, the exploration of novel applications and protection of the environment.

References 1. 2. 3. 4. 5. 6.

Woese CR, Kandler O, Wheelis ML (1990) Proc Natl Acad Sci USA 87:4576 Woese CR, Achenbach L, Rouviere P, Mandelco L (1991) Syst Appl Microbiol 14:364 Lorenz P, Liebeton K, Niehaus F, Eck J (2002) Curr Opin Biotechnol 13:572 Kauffmann IM, Schmitt J, Schmid RD (2004) Appl Microbiol Biotechnol 64:665 Beja O (2004) Curr Opin Biotechnol 15:187 Beja O, Suzuki MT, Heidelberg JF, Nelson WC, Preston CM, Hamada T, Eisen JA, Fraser CM, DeLong EF (2002) Nature 415:630 7. Beja O, Koonin EV, Aravind L, Taylor LT, Seitz H, Stein JL, Bensen DC, Feldman RA, Swanson RV, DeLong EF (2002) Appl Environ Microbiol 68:335 8. Beja O, Suzuki MT, Koonin EV, Aravind L, Hadd A, Nguyen LP, Villacorta R, Amjadi M, Garrigues C, Jovanovich SB, Feldman RA, DeLong EF (2000) Environ Microbiol 2:516 9. Beja O, Aravind L, Koonin EV, Suzuki MT, Hadd A, Nguyen LP, Jovanovich SB, Gates CM, Feldman RA, Spudich JL, Spudich EN, DeLong EF (2000) Science 289:1902

254

G. Antranikian et al.

10. Henne A, Schmitz RA, Bomeke M, Gottschalk G, Daniel R (2000) Appl Environ Microbiol 66:3113 11. Healy FG, Ray RM, Aldrich HC, Wilkie AC, Ingram LO, Shanmugam KT (1995) Appl Microbiol Biotechnol 43:667 12. Richardson TH, Tan X, Frey G, Callen W, Cabell M, Lam D, Macomber J, Short JM, Robertson DE, Miller C (2002) J Biol Chem 277:26 501 13. Cottrell MT, Moore JA, Kirchman DL (1999) Appl Environ Microbiol 65:2553 14. Cottrell MT, Wood DN, Yu L, Kirchman DL (2000) Appl Environ Microbiol 66:1195 15. Rondon MR, Raffel SJ, Goodman RM, Handelsman J (1999) Proc Natl Acad Sci USA 96:6451 16. Rondon MR, August PR, Bettermann AD, Brady SF, Grossman TH, Liles MR, Loiacono KA, Lynch BA, MacNeil IA, Minor C, Tiong CL, Gilman M, Osburne MS, Clardy J, Handelsman J, Goodman RM (2000) Appl Environ Microbiol 66:2541 17. Torsvik V, Ovreas L (2002) Curr Opin Microbiol 5:240 18. Torsvik V, Ovreas L, Thingstad TF (2002) Science 296:1064 19. Entcheva P, Liebl W, Johann A, Hartsch T, Streit WR (2001) Appl Environ Microbiol 67:89 20. Voget S, Leggewie C, Uesbeck A, Raasch C, Jaeger KE, Streit WR (2003) Appl Environ Microbiol 69:6235 21. Lorenz P, Scleper C (2002) J Mol Catal B 19–20:13 22. Morita RY (1975) Bacteriol Rev 39:144 23. Gerday C, Aittaleb M, Bentahir M, Chessa JP, Claverie P, Collins T, D’Amico S, Dumont J, Garsoux G, Georlette D, Hoyoux A, Lonhienne T, Meuwis MA, Feller G (2000) Trends Biotechnol 18:103 24. Lonhienne T, Gerday C, Feller G (2000) Biochim Biophys Acta 1543:1 25. Stetter KO (1996) Ciba Found Symp 202:1 26. Huber R, Huber H, Stetter KO (2000) FEMS Microbiol Rev 24:615 27. Segerer AH, Burggraf S, Fiala G, Huber G, Huber R, Pley U, Stetter KO (1993) Orig Life Evol Biosph 23:77 28. Stetter KO (1999) FEBS Lett 452:22 29. Lien T, Madsen M, Rainey FA, Birkeland NK (1998) Int J Syst Bacteriol 48 Pt 3:1007 30. Woese CR, Fox GE (1977) Proc Natl Acad Sci USA 74:5088 31. Jan RL, Wu J, Chaw SM, Tsai CW, Tsen SD (1999) Int J Syst Bacteriol 49 Pt 4:1809 32. Xiang X, Dong X, Huang L (2003) Extremophiles 7:493 33. He ZG, Zhong H, Li Y (2004) Curr Microbiol 48:159 34. Searcy DG, Stein DB (1980) Biochim Biophys Acta 609:180 35. Schleper C, Puehler G, Holz I, Gambacorta A, Janekovic D, Santarius U, Klenk HP, Zillig W (1995) J Bacteriol 177:7050 36. Smith PF (1980) Biochim Biophys Acta 619:367 37. van de Vossenberg, Driessen AJ, Zillig W, Konings WN (1998) Extremophiles 2:67 38. Futterer O, Angelov A, Liesegang H, Gottschalk G, Schleper C, Schepers B, Dock C, Antranikian G, Liebl W (2004) Proc Natl Acad Sci USA 101:9091 39. Bandeiras TM, Salgueiro CA, Huber H, Gomes CM, Teixeira M (2003) Biochim Biophys Acta 1557:13 40. Nemati M, Lowenadler J, Harrison ST (2000) Appl Microbiol Biotechnol 53:173 41. Burton NP, Williams TD, Norris PR (1999) Arch Microbiol 172:349 42. Gomes CM, Huber H, Stetter KO, Teixeira M (1998) FEBS Lett 432:99 43. Brierley CL, Brierley JA (1973) Can J Microbiol 19:183 44. Kurosawa N, Itoh YH, Itoh T (2003) Int J Syst Evol Microbiol 53:1607 45. Krulwich TA, Guffanti AA, Ito M (1999) Novartis Found Symp 221:167

Extreme Environments as a Resource for Microorganisms and Novel Biocatalysts 46. 47. 48. 49. 50. 51. 52. 53. 54. 55. 56. 57. 58. 59. 60. 61. 62. 63. 64. 65.

66. 67. 68. 69. 70. 71. 72. 73. 74. 75. 76. 77. 78. 79. 80. 81.

255

Horikoshi K (1999) Microbiol Mol Biol Rev 63:735 Jones BE, Grant WD, Duckworth AW, Owenson GG (1998) Extremophiles 2:191 Wiegel J, Kevbrin VV (2004) Biochem Soc Trans 32:193 Prowe SG, Antranikian G (2001) Int J Syst Evol Microbiol 51:457 Engle M, Li Y, Woese C, Wiegel J (1995) Int J Syst Bacteriol 45:454 Keller M, Braun FJ, Dirmeier R, Hafenbradl D, Burggraf S, Rachel R, Stetter KO (1995) Arch Microbiol 164:390 Dirmeier R, Keller M, Hafenbradl D, Braun FJ, Rachel R, Burggraf S, Stetter KO (1998) Extremophiles 2:109 Litchfield CD, Gillevet PM (2002) J Ind Microbiol Biotechnol 28:48 Sanchez-Porro C, Martin S, Mellado E, Ventosa A (2003) J Appl Microbiol 94:295 Ventosa A, Marquez MC, Garabito MJ, Arahal DR (1998) Extremophiles 2:297 Ollivier B, Caumette P, Garcia JL, Mah RA (1994) Microbiol Rev 58:27 Ventosa A, Nieto JJ, Oren A (1998) Microbiol Mol Biol Rev 62:504 Liebl W (2001) Methods Enzymol 330:290 Bronnenmeier K, Kern A, Liebl W, Staudenbauer WL (1995) Appl Environ Microbiol 61:1399 Bok JD, Yernool DA, Eveleigh DE (1998) Appl Environ Microbiol 64:4774 Te’o VS, Saul DJ, Bergquist PL (1995) Appl Microbiol Biotechnol 43:291 Saul DJ, Williams LC, Grayling RA, Chamley LW, Love DR, Bergquist PL (1990) Appl Environ Microbiol 56:3117 Saul DJ, Williams LC, Love DR, Chamley LW, Bergquist PL (1989) Nucleic Acids Res 17:439 Ruttersmith LD, Daniel RM (1991) Biochem J 277 (Pt 3):887 Halldorsdottir S, Thorolfsdottir ET, Spilliaert R, Johansson M, Thorbjarnardottir SH, Palsdottir A, Hreggvidsson GO, Kristjansson JK, Holst O, Eggertsson G (1998) Appl Microbiol Biotechnol 49:277 Crennell SJ, Hreggvidsson GO, Nordberg Karlsson E (2002) J Mol Biol 320:883 Bauer MW, Driskill LE, Callen W, Snead MA, Mathur EJ, Kelly RM (1999) J Bacteriol 181:284 Moracci M, Ciaramella M, Rossi M (2001) Methods Enzymol 330:201 Bauer MW, Bylina EJ, Swanson RV, Kelly RM (1996) J Biol Chem 271:23 749 Bauer MW, Kelly RM (1998) Biochemistry 37:17 170 Nucci R, Moracci M, Vaccaro C, Vespa N, Rossi M (1993) Biotechnol Appl Biochem 17 (Pt 2):239 Moracci M, Nucci R, Febbraio F, Vaccaro C, Vespa N, La Cara F, Rossi M (1995) Enzyme Microb Technol 17:992 Sivori AS, Mercuri OA, Forchiassin F (1996) Rev Argent Microbiol 28:9 Durrant LR, Mello AB, Reginatto V (1992) Biochem Soc Trans 20:227S Allen WG (1976) Biotechnol Bioeng Symp 6:303 Schulein M (2000) Biochim Biophys Acta 1543:239 Jeffries TW (1996) Curr Opin Biotechnol 7:337 Te’o VS, Cziferszky AE, Bergquist PL, Nevalainen KM (2000) FEMS Microbiol Lett 190:13 McCarthy AA, Morris DD, Bergquist PL, Baker EN (2000) Acta Crystallogr D Biol Crystallogr 56 (Pt 11):1367 Simpson HD, Haufler UR, Daniel RM (1991) Biochem J 277 (Pt 2):413 Winterhalter C, Heinrich P, Candussio A, Wich G, Liebl W (1995) Mol Microbiol 15:431

256

G. Antranikian et al.

82. Velikodvorskaya TV, Volkov I, Vasilevko VT, Zverlov VV, Piruzian ES (1997) Biochemistry (Mosc) 62:66 83. Zverlov V, Piotukh K, Dakhova O, Velikodvorskaya G, Borriss R (1996) Appl Microbiol Biotechnol 45:245 84. Walsh DJ, Gibbs MD, Bergquist PL (1998) Extremophiles 2:9 85. Meissner K, Wassenberg D, Liebl W (2000) Mol Microbiol 36:898 86. Lama L, Calandrelli V, Gambacorta A, Nicolaus B (2004) Res Microbiol 155:283 87. Teplitsky A, Mechaly A, Stojanoff V, Sainz G, Golan G, Feinberg H, Gilboa R, Reiland V, Zolotnitsky G, Shallom D, Thompson A, Shoham Y, Shoham G (2004) Acta Crystallogr D Biol Crystallogr 60:836 88. Andrade CM, Aguiar WB, Antranikian G (2001) Appl Biochem Biotechnol 91-93:655 89. Waino M, Ingvorsen K (2003) Extremophiles 7:87 90. Linko M, Viikari L, Suihko ML (1984) Biotechnol Adv 2:233 91. Oksanen T, Pere J, Paavilainen L, Buchert J, Viikari L (2000) J Biotechnol 78:39 92. Schink B, Zeikus JB (1983) FEMS Microbiol Lett 17:295 93. Tamaru Y, Doi RH (2001) Proc Natl Acad Sci USA 98:4125 94. Van Rijssel M, Gerwig GJ, Hansen TA (1993) Appl Environ Microbiol 59:828 95. Hoondal GS, Tiwari RP, Tewari R, Dahiya N, Beg QK (2002) Appl Microbiol Biotechnol 59:409 96. Kashyap DR, Vohra PK, Chopra S, Tewari R (2001) Bioresour Technol 77:215 97. Herron SR, Benen JA, Scavetta RD, Visser J, Jurnak F (2000) Proc Natl Acad Sci USA 97:8762 98. Prade RA, Zhan D, Ayoubi P, Mort AJ (1999) Biotechnol Genet Eng Rev 16:361 99. Lang C, Dornenburg H (2000) Appl Microbiol Biotechnol 53:366 100. Sakai T, Sakamoto T, Hallaert J, Vandamme EJ (1993) Adv Appl Microbiol 39:213 101. Rexova-Benkova L, Markovic O (1976) Adv Carbohydr Chem Biochem 33:323 102. Kozianowski G, Canganella F, Rainey FA, Hippe H, Antranikian G (1997) Extremophiles 1:171 103. Kluskens LD, van Alebeek GJ, Voragen AG, de Vos WM, van der Oost J (2003) Biochem J 370:651 104. Cohen-Kupiec R, Chet I (1998) Curr Opin Biotechnol 9:270 105. Koga D, Mitsutomi M, Kono M, Matsumiya M (1999) EXS 87:111 106. Fukamizo T (2000) Curr Protein Pept Sci 1:105 107. Takayanagi T, Ajisaka K, Takiguchi Y, Shimahara K (1991) Biochim Biophys Acta 1078:404 108. Sakai K, Yokota A, Kurokawa H, Wakayama M, Moriguchi M (1998) Appl Environ Microbiol 64:3397 109. Tsujibo H, Minoura K, Miyamoto K, Endo H, Moriwaki M, Inamori Y (1993) Appl Environ Microbiol 59:620 110. Andronopoulou E, Vorgias CE (2004) Protein Expr Purif 35:264 111. Imanaka T, Fukui T, Fujiwara S (2001) Methods Enzymol 330:319 112. Tanaka T, Fujiwara S, Nishikori S, Fukui T, Takagi M, Imanaka T (1999) Appl Environ Microbiol 65:5338 113. Gao J, Bauer MW, Shockley KR, Pysz MA, Kelly RM (2003) Appl Environ Microbiol 69:3119 114. Grzybowska B, Szweda P, Synowiecki J (2004) Mol Biotechnol 26:101 115. Berk H, Lebbink RJ (2003) Methods Mol Biol 230:127 116. Linden A, Mayans O, Meyer-Klaucke W, Antranikian G, Wilmanns M (2003) J Biol Chem 278:9875 117. Savchenko A, Vieille C, Kang S, Zeikus JG (2002) Biochemistry 41:6193

Extreme Environments as a Resource for Microorganisms and Novel Biocatalysts 118. 119. 120. 121. 122. 123. 124. 125. 126. 127. 128. 129. 130. 131. 132. 133. 134. 135. 136. 137. 138. 139. 140. 141. 142. 143. 144. 145. 146. 147. 148. 149. 150. 151. 152. 153. 154. 155.

257

Savchenko A, Vieille C, Zeikus JG (2001) Methods Enzymol 330:354 Leveque E, Haye B, Belarbi A (2000) FEMS Microbiol Lett 186:67 Canganella F, Gambacorta A, Kato C, Horikoshi K (2000) Microbiol Res 154:297 Kato M, Miura Y, Kettoku M, Shindo K, Iwamatsu A, Kobayashi K (1996) Biosci Biotechnol Biochem 60:546 Kobayashi K, Kato M, Miura Y, Kettoku M, Komeda T, Iwamatsu A (1996) Biosci Biotechnol Biochem 60:1720 Haseltine C, Rolfsmeier M, Blum P (1996) J Bacteriol 178:945 Lee J, Kanai H, Kobayashi T, Akiba T, Kudo T (1996) J Ferment Bioeng 82:432 Tachibana Y, Mendez L, Fujiwara S, Takagi M, Imanaka T (1996) J Ferment Bioeng 82:224 Liebl W, Stemplinger I, Ruile P (1997) J Bacteriol 179:941 Horinouchi S, Fukusumi S, Ohshima T, Beppu T (1988) Eur J Biochem 176:243 Fukusumi S, Kamizono A, Horinouchi S, Beppu T (1988) Eur J Biochem 174:15 Ducki A, Grundmann O, Konermann L, Mayer F, Hoppert M (1998) J Gen Appl Microbiol 44:327 Ganghofner D, Kellermann J, Staudenbauer WL, Bronnenmeier K (1998) Biosci Biotechnol Biochem 62:302 Coutinho PM, Reilly PJ (1994) Protein Eng 7:749 Coutinho PM, Reilly PJ (1994) Protein Eng 7:393 Ohnishi H, Matsumoto H, Sakai H, Ohta T (1994) J Biol Chem 269:3503 Ohnishi H, Kitamura H, Minowa T, Sakai H, Ohta T (1992) Eur J Biochem 207:413 Paquet V, Croux C, Goma G, Soucaille P (1991) Appl Environ Microbiol 57:212 Glushchenko EV, Kozlov DG, Podkovyrov SM, Mogutov MA (1990) Mol Biol (Mosk) 24:744 Kozlov DG, Kurepina NE, Glushchenko EV, Mogutov MA, Podkovyrov SM (1990) Mol Biol (Mosk) 24:736 Saha BC, Mathupala SP, Zeikus JG (1988) Biochem J 252:343 Hyun HH, Shen GJ, Zeikus JG (1985) J Bacteriol 164:1153 Hyun HH, Zeikus JG (1985) J Bacteriol 164:1146 Hyun HH, Zeikus JG (1985) J Bacteriol 164:1162 Serour E, Antranikian G (2002) Antonie Van Leeuwenhoek 81:73 Kim MS, Park JT, Kim YW, Lee HS, Nyawira R, Shin HS, Park CS, Yoo SH, Kim YR, Moon TW, Park KH (2004) Appl Environ Microbiol 70:3933 Chang ST, Parker KN, Bauer MW, Kelly RM (2001) Methods Enzymol 330:260 Galichet A, Belarbi A (1999) FEBS Lett 458:188 Xavier KB, Peist R, Kossmann M, Boos W, Santos H (1999) J Bacteriol 181:3358 Schicho RN, Ma K, Adams MW, Kelly RM (1993) J Bacteriol 175:1823 Costantino HR, Brown SH, Kelly RM (1990) J Bacteriol 172:3654 Miura Y, Kettoku M, Kato M, Kobayashi K, Kondo K (1999) J Mol Microbiol Biotechnol 1:129 Campbell BS, Siddique AB, McDougall BM, Seviour RJ (2004) FEMS Microbiol Lett 232:225 Bertoldo C, Antranikian G (2002) Curr Opin Chem Biol 6:151 Bertoldo C, Antranikian G (2001) Methods Enzymol 330:269 Dong G, Vieille C, Zeikus JG (1997) Appl Environ Microbiol 63:3577 Canganella F, Jones WJ, Gambacorta A, Antranikian G (1997) Arch Microbiol 167:233 Duffner F, Bertoldo C, Andersen JT, Wagner K, Antranikian G (2000) J Bacteriol 182:6331

258

G. Antranikian et al.

156. Erra-Pujada M, Debeire P, Duchiron F, O’Donohue MJ (1999) J Bacteriol 181:3284 157. Niehaus F, Peters A, Groudieva T, Antranikian G (2000) FEMS Microbiol Lett 190:223 158. Knapp S, Rudiger A, Antranikian G, Jorgensen PL, Ladenstein R (1995) Proteins 23:595 159. Rudiger A, Jorgensen PL, Antranikian G (1995) Appl Environ Microbiol 61:567 160. Schwerdtfeger RM, Chiaraluce R, Consalvi V, Scandurra R, Antranikian G (1999) Eur J Biochem 264:479 161. Zona R, Chang-Pi-Hin F, O’Donohue MJ, Janecek S (2004) Eur J Biochem 271:2863 162. Kang S, Vieille C, Zeikus JG (2005) Appl Microbiol Biotechnol 66:408 163. Kim CH, Nashiru O, Ko JH (1996) FEMS Microbiol Lett 138:147 164. Bertoldo C, Duffner F, Jorgensen PL, Antranikian G (1999) Appl Environ Microbiol 65:2084 165. Lebbink JH, Bertoldo C, Tibbelin G, Andersen JT, Duffner F, Antranikian G, Ladenstein R (2000) Acta Crystallogr D Biol Crystallogr 56 (Pt 11):1470 166. Koch R, Canganella F, Hippe H, Jahnke K, Antranikian G (1997) Appl Environ Microbiol 63:1088 167. Bibel M, Brettl C, Gosslar U, Kriegshauser G, Liebl W (1998) FEMS Microbiol Lett 158:9 168. Bertoldo C, Armbrecht M, Becker F, Schafer T, Antranikian G, Liebl W (2004) Appl Environ Microbiol 70:3407 169. Wind RD, Liebl W, Buitelaar RM, Penninga D, Spreinat A, Dijkhuizen L, Bahl H (1995) Appl Environ Microbiol 61:1257 170. Knegtel RM, Wind RD, Rozeboom HJ, Kalk KH, Buitelaar RM, Dijkhuizen L, Dijkstra BW (1996) J Mol Biol 256:611 171. Tachibana Y, Kuramura A, Shirasaka N, Suzuki Y, Yamamoto T, Fujiwara S, Takagi M, Imanaka T (1999) Appl Environ Microbiol 65:1991 172. Biwer A, Antranikian G, Heinzle E (2002) Appl Microbiol Biotechnol 59:609 173. Cowan DA, Smolenski KA, Daniel RM, Morgan HW (1987) Biochem J 247:121 174. Guagliardi A, Cerchia L, Rossi M (2002) Biochem J 368:357 175. Burlini N, Magnani P, Villa A, Macchi F, Tortora P, Guerritore A (1992) Biochim Biophys Acta 1122:283 176. Mayr J, Lupas A, Kellermann J, Eckerskorn C, Baumeister W, Peters J (1996) Curr Biol 6:739 177. Kannan Y, Koga Y, Inoue Y, Haruki M, Takagi M, Imanaka T, Morikawa M, Kanaya S (2001) Appl Environ Microbiol 67:2445 178. Morikawa M, Imanaka T (2001) Methods Enzymol 330:424 179. Volkl P, Markiewicz P, Stetter KO, Miller JH (1994) Protein Sci 3:1329 180. Chang LS, Hicks PM, Kelly RM (2001) Methods Enzymol 330:403 181. Morikawa M, Izawa Y, Rashid N, Hoaki T, Imanaka T (1994) Appl Environ Microbiol 60:4559 182. Klingeberg M, Galunsky B, Sjoholm C, Kasche V, Antranikian G (1995) Appl Environ Microbiol 61:3098 183. Chavez Croocker P, Sako Y, Uchida A (1999) Extremophiles 3:3 184. Sako Y, Croocker PC, Ishida Y (1997) FEBS Lett 415:329 185. Catara G, Ruggiero G, La Cara F, Digilio FA, Capasso A, Rossi M (2003) Extremophiles 7:391 186. Voorhorst WG, Eggen RI, Geerling AC, Platteeuw C, Siezen RJ, Vos WM (1996) J Biol Chem 271:20 426 187. de Vos WM, Voorhorst WG, Dijkgraaf M, Kluskens LD, Van der Oost J, Siezen RJ (2001) Methods Enzymol 330:383

Extreme Environments as a Resource for Microorganisms and Novel Biocatalysts 188. 189. 190. 191. 192. 193. 194. 195. 196. 197. 198. 199. 200. 201. 202. 203. 204. 205. 206. 207. 208. 209. 210. 211. 212. 213. 214. 215. 216. 217. 218. 219. 220. 221. 222. 223. 224. 225. 226.

259

Voorhorst WG, Warner A, de Vos WM, Siezen RJ (1997) Protein Eng 10:905 Choi IG, Bang WG, Kim SH, Yu YG (1999) J Biol Chem 274:881 Lin X, Fusek M, Tang J (1991) Adv Exp Med Biol 306:255 Fusek M, Lin XL, Tang J (1990) J Biol Chem 265:1496 Lin X, Tang J (1990) J Biol Chem 265:1490 Fujiwara S, Okuyama S, Imanaka T (1996) Gene 179:165 Friedrich A, Antranikian G (1996) Appl Environ Microbiol 62:2875 Kim JS, Kluskens LD, de Vos WM, Huber R, van der Oost J (2004) J Mol Biol 335:787 Kluskens LD, Voorhorst WG, Siezen RJ, Schwerdtfeger RM, Antranikian G, van der Oost J, de Vos WM (2002) Extremophiles 6:185 Lesuisse E, Schanck K, Colson C (1993) Eur J Biochem 216:155 Sinchaikul S, Sookkheo B, Phutrakul S, Pan FM, Chen ST (2001) Protein Expr Purif 22:388 Cho AR, Yoo SK, Kim EJ (2000) FEMS Microbiol Lett 186:235 Lee D, Koh Y, Kim K, Kim B, Choi H, Kim D, Suhartono MT, Pyun Y (1999) FEMS Microbiol Lett 179:393 Tyndall JD, Sinchaikul S, Fothergill-Gilmore LA, Taylor P, Walkinshaw MD (2002) J Mol Biol 323:859 Crabb WD, Shetty JK (1999) Curr Opin Microbiol 2:252 Park BC, Koh S, Chang C, Suh SW, Lee DS, Byun SM (1997) Appl Biochem Biotechnol 62:15 Liu SY, Wiegel J, Gherardini FC (1996) J Bacteriol 178:5938 Vieille C, Hess JM, Kelly RM, Zeikus JG (1995) Appl Environ Microbiol 61:1867 Vieille C, Sriprapundh D, Kelly RM, Zeikus JG (2001) Methods Enzymol 330:215 Hess JM, Kelly RM (1999) Biotechnol Bioeng 62:509 Hess JM, Tchernajenko V, Vieille C, Zeikus JG, Kelly RM (1998) Appl Environ Microbiol 64:2357 Cowan D (1996) Trends Biotech 14:177 Burdette DS, Vieille C, Zeikus JG (1996) Biochem J 316 (Pt 1):115 Burdette DS, Jung SH, Shen GJ, Hollingsworth RI, Zeikus JG (2002) Appl Environ Microbiol 68:1914 Burdette DS, Tchernajencko VV, Zeikus JG (2000) Enzyme Microb Technol 27:11 Burdette DS, Secundo F, Phillips RS, Dong J, Scott RA, Zeikus JG (1997) Biochem J 326 (Pt 3):717 Yoon SY, Noh HS, Kim EH, Kong KH (2002) Comp Biochem Physiol B Biochem Mol Biol 132:415 Kong KH, Hong MP, Choi SS, Kim YT, Cho SH (2000) Biotechnol Appl Biochem 31 (Pt 2):113 Abokitse K, Hummel W (2003) Appl Microbiol Biotechnol 62:380 Ma K, Adams MW (2001) Methods Enzymol 331:195 Li D, Stevenson KJ (2001) Methods Enzymol 331:201 Zhang J, Liu J, Zhou J, Ren Y, Dai X, Xiang H (2003) Biotechnol Lett 25:1463 Suzuki Y, Miyamoto K, Ohta H (2004) FEMS Microbiol Lett 236:97 Ikeda M, Clark DS (1998) Biotechnol Bioeng 57:624 Labahn J, Neumann S, Buldt G, Kula MR, Granzin J (2002) J Mol Biol 322:1053 Chebrou H, Bigey F, Arnaud A, Galzy P (1996) Biochim Biophys Acta 1298:285 Kobayashi M, Komeda H, Nagasawa T, Nishiyama M, Horinouchi S, Beppu T, Yamada H, Shimizu S (1993) Eur J Biochem 217:327 Ciskanik l, Wilczek JM, Fallon RD (1995) Appl Environ Microbiol 61:998 Trott S, Bauer R, Knackmuss HJ, Stolz A (2001) Microbiology 147:1815

260

G. Antranikian et al.

227. Mayaux J, Cerbelaud E, Soubrier F, Yeh P, Blanche F, Petre D (1991) J Bacteriol 173:6694 228. Hirrlinger B, Stolz A, Knackmuss HJ (1996) J Bacteriol 178:3501 229. Layh N, Hirrlinger B, Stolz A, Knackmuss HJ (1997) Appl Microbiol Biotechnol 47:668 230. Hermes H, Tandler RF, Sonke T, Dijkhuizen L, Meijer EM (1994) Appl Environ Microbiol 1:153 231. Sonke T, Kaptein B, Boesten WHJ, Boxterman QB, Schoemaker HE, Kamphuis J, Formaggio F, Toniolo C, Rutjes FP (2000) In: RN P (ed) Stereoselective Biocatalysis. Marcel Dekker, New York p 23 232. Kamphuis J, Boesten WHJ, Kaptein B, Hermes HFM, Sonke T, Broxterman QB, van den Tweel WJJ, Schoemaker HE (1992) in: Collins AN, Sheldrake GN, Crosby J (eds) Chirality in industry: the commercial manufacture and applications of optically active compounds. Wiley, Chichester, UK p 187 233. Alcantara A-R, Sanchez-Montero JM, Sinisterra JV (2000) In: RN P (ed) Stereoselective biocatalysis. Marcel Dekker, New York p 659 234. Webster N, Ramsden DK, Hughes J (2001) Biotechnol Lett 23:95 235. Fournand D, Arnaud A, Galzy P (1998) J Mol Catal 4:77 236. Baek DH, Song JJ, Lee SG, Kwon SJ, Asano Y, Sung MH (2003) Enzyme Microb Technol 32:131 237. Nawaz M, Khan AA, Bhattacharayya D, Siitonen PH, Cerniglia CE (1996) J Bacteriol 178:2397 238. d’Abusco AS, Ammendola S, Scandurra R, Politi L (2001) Extremophiles 5:183 239. Egorova K, Trauthwein H, Verseck S, Antranikian G (2004) Appl Microbiol Biotechnol 65:38 240. Mahadevan S, Thimann KV (1964) Arch Biochem Biophys 107:62 241. Thimann KV, Mahadevan S (1964) Arch Biochem Biophys 105:133 242. Hook RH, Robinson WG (1964) J Biol Chem 239:4263 243. Robinson WG, Hook RH (1964) J Biol Chem 239:4257 244. Arnaud A, Galzy P, Jallageas JC (1976) C R Acad Sci Hebd Seances Acad Sci D 283:571 245. Harper DB (1977) Biochem J 167:685 246. Harper DB (1977) Biochem J 165:309 247. Wajant H, Effenberger F (2002) Eur J Biochem 269:680 248. Pekarsky Y, Campiglio M, Siprashvili Z, Druck T, Sedkov Y, Tillib S, Draganescu A, Wermuth P, Rothman JH, Huebner K, Buchberg AM, Mazo A, Brenner C, Croce CM (1998) Proc Natl Acad Sci USA 95:8744 249. Pace HC, Hodawadekar SC, Draganescu A, Huang J, Bieganowski P, Pekarsky Y, Croce CM, Brenner C (2000) Curr Biol 10:907 250. Harper DB (1985) Int J Biochem 17:677 251. Stevenson DE, Feng R, Dumas F, Groleau D, Mihoc A, Storer AC (1992) Biotechnol Appl Biochem 15:283 252. Nagasawa T, Mauger J, Yamada H (1990) Eur J Biochem 194:765 253. Kobayashi M, Komeda H, Yanaka N, Nagasawa T, Yamada H (1992) J Biol Chem 267:20 746 254. Kobayashi M, Yanaka N, Nagasawa T, Yamada H (1992) Biochemistry 31:9000 255. Banerjee A, Sharma R, Banerjee UC (2002) Appl Microbiol Biotechnol 60:33 256. Perler FB, Kumar S, Kong H (1996) Adv Protein Chem 48:377 257. Mullis K, Faloona F, Scharf S, Saiki R, Horn G, Erlich H (1992) Biotechnology 24:17

Extreme Environments as a Resource for Microorganisms and Novel Biocatalysts

261

258. Saiki RK, Gelfand DH, Stoffel S, Scharf SJ, Higuchi R, Horn GT, Mullis KB, Erlich HA (1988) Science 239:487 259. Mullis K, Faloona F, Scharf S, Saiki R, Horn G, Erlich H (1986) Cold Spring Harb Symp Quant Biol 51 Pt 1:263 260. Diaz RS, Sabino EC (1998) Braz J Med Biol Res 31:1239 261. Komori K, Ishino Y (2000) Protein Eng 13:41 262. Evans SJ, Fogg MJ, Mamone A, Davis M, Pearl LH, Connolly BA (2000) Nucleic Acids Res 28:1059 263. Hayashi I, Morikawa K, Ishino Y (1999) Nucleic Acids Res 27:4695 264. Goldman S, Kim R, Hung LW, Jancarik J, Kim SH (1998) Acta Crystallogr D Biol Crystallogr 54 (Pt 5):986 265. Lu C, Erickson HP (1997) Protein Expr Purif 11:179 266. Uemori T, Sato Y, Kato I, Doi H, Ishino Y (1997) Genes Cells 2:499 267. Imamura M, Uemori T, Kato I, Ishino Y (1995) Biol Pharm Bull 18:1647 268. Uemori T, Ishino Y, Toh H, Asada K, Kato I (1993) Nucleic Acids Res 21:259 269. Ogata N, Miura T (2000) Biochemistry 39:13 993 270. Ogata N, Miura T (1998) Nucleic Acids Res 26:4652 271. Keohavong P, Ling L, Dias C, Thilly WG (1993) PCR Methods Appl 2:288 272. Kong H, Kucera RB, Jack WE (1993) J Biol Chem 268:1965 273. Mattila P, Korpela J, Tenkanen T, Pitkanen K (1991) Nucleic Acids Res 19:4967 274. Cariello NF, Swenberg JA, Skopek TR (1991) Nucleic Acids Res 19:4193 275. Hashimoto H, Nishioka M, Fujiwara S, Takagi M, Imanaka T, Inoue T, Kai Y (2001) J Mol Biol 306:469 276. Hashimoto H, Matsumoto T, Nishioka M, Yuasa T, Takeuchi S, Inoue T, Fujiwara S, Takagi M, Imanaka T, Kai Y (1999) J Biochem (Tokyo) 125:983 277. Nishioka M, Fujiwara S, Takagi M, Imanaka T (1998) Nucleic Acids Res 26:4409 278. Takagi M, Nishioka M, Kakihara H, Kitabayashi M, Inoue H, Kawakami B, Oka M, Imanaka T (1997) Appl Environ Microbiol 63:4504 279. Sanger F, Nicklen S, Coulson AR (1992) Biotechnology 24:104 280. Bechtereva TA, Pavlov YI, Kramorov VI, Migunova B, Kiselev OI (1989) Nucleic Acids Res 17:10 507 281. Rao VB, Saunders NB (1992) Gene 113:17 282. Vander Horn PB, Davis MC, Cunniff JJ, Ruan C, McArdle BF, Samols SB, Szasz J, Hu G, Hujer KM, Domke ST, Brummet SR, Moffett RB, Fuller CW (1997) Biotechniques 22:758 283. Szybalski W (1990) Gene 90:177 284. Schalling M, Hudson TJ, Buetow KH, Housman DE (1993) Nat Genet 4:135 285. Landegren U, Schallmeiner E, Nilsson M, Fredriksson S, Baner J, Gullberg M, Jarvius J, Gustafsdottir S, Dahl F, Soderberg O, Ericsson O, Stenberg J (2004) J Mol Recognit 17:194 286. Georlette D, Blaise V, Bouillenne F, Damien B, Thorbjarnardottir SH, Depiereux E, Gerday C, Uversky VN, Feller G (2004) Biophys J 86:1089 287. Housby JN, Southern EM (2002) Anal Biochem 302:88 288. Jonsson ZO, Thorbjarnardottir SH, Eggertsson G, Palsdottir A (1994) Gene 151:177 289. Thorbjarnardottir SH, Jonsson ZO, Andresson OS, Kristjansson JK, Eggertsson G, Palsdottir A (1995) Gene 161:1 290. Blondel T, Hjorleifsdottir SH, Fridjonsson OF, Aevarsson A, Skirnisdottir S, Hermannsdottir AG, Hreggvidsson GO, Smith AV, Kristjansson JK (2003) Nucleic Acid Res 31:7247

262

G. Antranikian et al.

291. Kletzin A (1992) Nucleic Acid Res 20:5389 292. Nakatani M, Ezaki S, Atomi H, Imanaka T (2002) Eur J Biochem 269:650 293. Rolland JL, Guenguen Y, Persillon C, Masson JM, Dietrich J (2004) FEMS Microbiol Lett 236:267

Author Index Volumes 51–97 Author Index Volumes 1–50 see Volume 50

Ackermann, J.-U. see Babel, W.: Vol. 71, p. 125 Adam, W., Lazarus, M., Saha-Möller, C. R., Weichhold, O., Hoch, U., Häring, D., Schreier, Ü.: Biotransformations with Peroxidases. Vol. 63, p. 73 Ahring, B. K.: Perspectives for Anaerobic Digestion. Vol. 81, p. 1 Ahring, B. K. see Angelidaki, I.: Vol. 82, p. 1 Ahring, B. K. see Gavala, H. N.: Vol. 81, p. 57 Ahring, B. K. see Hofman-Bang, J.: Vol. 81, p. 151 Ahring, B. K. see Mogensen, A. S.: Vol. 82, p. 69 Ahring, B. K. see Pind, P. F.: Vol. 82, p. 135 Ahring, B. K. see Skiadas, I. V.: Vol. 82, p. 35 Aivasidis, A., Diamantis, V. I.: Biochemical Reaction Engineering and Process Development in Anaerobic Wastewater Treatment. Vol. 92, p. 49 Akhtar, M., Blanchette, R. A., Kirk, T. K.: Fungal Delignification and Biochemical Pulping of Wood. Vol. 57, p. 159 Allan, J. V., Roberts, S. M., Williamson, N. M.: Polyamino Acids as Man-Made Catalysts. Vol. 63, p. 125 Allington, R. W. see Xie, S.: Vol. 76, p. 87 Al-Abdallah, Q. see Brakhage, A. A.: Vol. 88, p. 45 Al-Rubeai, M.: Apoptosis and Cell Culture Technology. Vol. 59, p. 225 Al-Rubeai, M. see Singh, R. P.: Vol. 62, p. 167 Alsberg, B. K. see Shaw, A. D.: Vol. 66, p. 83 Angelidaki, I., Ellegaard, L., Ahring, B. K.: Applications of the Anaerobic Digestion Process. Vol. 82, p. 1 Angelidaki, I. see Gavala, H. N.: Vol. 81, p. 57 Angelidaki, I. see Pind, P. F.: Vol. 82, p. 135 Antranikian, G. see Ladenstein, R.: Vol. 61, p. 37 Antranikian, G. see Müller, R.: Vol. 61, p. 155 Antranikian, G., Vorgias, C. E., Bertoldo, C.: Extreme Environments as a Resource for Microorganisms and Novel Biocatalysts. Vol. 96, p. 219 Archelas, A. see Orru, R. V. A.: Vol. 63, p. 145 Argyropoulos, D. S.: Lignin. Vol. 57, p. 127 Arnold, F. H., Moore, J. C.: Optimizing Industrial Enzymes by Directed Evolution. Vol. 58, p. 1 Atala, A.: Regeneration of Urologic Tissues and Organs. Vol. 94, p. 181 Autuori, F., Farrace, M. G., Oliverio, S., Piredda, L., Piacentini, G.: “Tissieâ” Transglutaminase and Apoptosis. Vol. 62, p. 129 Azerad, R.: Microbial Models for Drug Metabolism. Vol. 63, p. 169

264

Author Index Volumes 51–97

Babel, W., Ackermann, J.-U., Breuer, U.: Physiology, Regulation and Limits of the Synthesis of Poly(3HB). Vol. 71, p. 125 Bajpai, P., Bajpai, P. K.: Realities and Trends in Emzymatic Prebleaching of Kraft Pulp. Vol. 56, p. 1 Bajpai, P., Bajpai, P. K.: Reduction of Organochlorine Compounds in Bleach Plant Effluents. Vol. 57, p. 213 Bajpai, P. K. see Bajpai, P.: Vol. 56, p. 1 Bajpai, P. K. see Bajpai, P.: Vol. 57, p. 213 Banks, M. K., Schwab, P., Liu, B., Kulakow, P. A., Smith, J. S., Kim, R.: The Effect of Plants on the Degradation and Toxicity of Petroleum Contaminants in Soil: A Field Assessment. Vol. 78, p. 75 Barber, M. S., Giesecke, U., Reichert, A., Minas, W.: Industrial Enzymatic Production of Cephalosporin-Based b-Lactams. Vol. 88, p. 179 Barindra, S. see Debashish, G.: Vol. 96, p. 189 Barnathan, G. see Bergé, J.-P.: Vol. 96, p. 49 Barut, M. see Strancar, A.: Vol. 76, p. 49 Bárzana, E.: Gas Phase Biosensors. Vol. 53, p. 1 Basu, S. K. see Mukhopadhyay, A.: Vol. 84, p. 183 Bathe, B. see Pfefferle, W.: Vol. 79, p. 59 Bazin, M. J. see Markov, S. A.: Vol. 52, p. 59 Bellgardt, K.-H.: Process Models for Production of b-Lactam Antibiotics. Vol. 60, p. 153 Beppu, T.: Development of Applied Microbiology to Modern Biotechnology in Japan. Vol. 69, p. 41 van den Berg, M. A. see Evers, M. E.: Vol. 88, p. 111 Bergé, J.-P., Barnathan, G.: Fatty Acids from Lipids of Marine Organisms: Molecular Biodiversity, Roles as Biomarkers, Biologically-Active Compounds, and Economical Aspects. Vol. 96, p. 49 Berovic, M. see Mitchell, D. A.: Vol. 68, p. 61 Bertoldo, C. see Antranikian, G.: Vol. 96, p. 219 Beyeler, W., DaPra, E., Schneider, K.: Automation of Industrial Bioprocesses. Vol. 70, p. 139 Beyer, M. see Seidel, G.: Vol. 66, p. 115 Beyer, M. see Tollnick, C.: Vol. 86, p. 1 Bhardwaj, D. see Chauhan, V. S.: Vol. 84, p. 143 Bhatia, P. K., Mukhopadhyay, A.: Protein Glycosylation: Implications for in vivo Functions and Thereapeutic Applications. Vol. 64, p. 155 Bisaria, V. S. see Ghose, T. K.: Vol. 69, p. 87 Blanchette R. A. see Akhtar, M.: Vol. 57, p. 159 Bocker, H., Knorre, W. A.: Antibiotica Research in Jena from Penicillin and Nourseothricin to Interferon. Vol. 70, p. 35 de Bont, J. A. M. see van der Werf, M. J.: Vol. 55, p. 147 van den Boom, D. see Jurinke, C.: Vol. 77, p. 57 Borah, M. M. see Dutta, M.: Vol. 86, p. 255 Bourguet-Kondracki, M.-L., Kornprobst, J.-M.: Marine Pharmacology: Potentialities in the Treatment of Infectious Diseases, Osteoporosis and Alzheimer’s Disease. Vol. 97, p. 105 Bovenberg, R. A. L. see Evers, M. E.: Vol. 88, p. 111 Brainard, A. P. see Ho, N. W. Y.: Vol. 65, p. 163 Brakhage, A. A., Spröte, P., Al-Abdallah, Q., Gehrke, A., Plattner, H., Tüncher, A.: Regulation of Penicillin Biosynthesis in Filamentous Fungi. Vol. 88, p. 45 Brazma, A., Sarkans, U., Robinson, A., Vilo, J., Vingron, M., Hoheisel, J., Fellenberg, K.: Microarray Data Representation, Annotation and Storage. Vol. 77, p. 113

Author Index Volumes 51–97

265

Breuer, U. see Babel, W.: Vol. 71, p. 125 Broadhurst, D. see Shaw, A. D.: Vol. 66, p. 83 Bruckheimer, E. M., Cho, S. H., Sarkiss, M., Herrmann, J., McDonell, T. J.: The Bcl-2 Gene Family and Apoptosis. Vol. 62, p. 75 Brüggemann, O.: Molecularly Imprinted Materials – Receptors More Durable than Nature Can Provide. Vol. 76, p. 127 Bruggink, A., Straathof, A. J. J., van der Wielen, L. A. M.: A ‘Fine’ Chemical Industry for Life Science Products: Green Solutions to Chemical Challenges. Vol. 80, p. 69 Buchert, J. see Suurnäkki, A.: Vol. 57, p. 261 Büchs, J. see Knoll, A.: Vol. 92, p. 77 Bungay, H. R. see Mühlemann, H. M.: Vol. 65, p. 193 Bungay, H. R., Isermann, H. P.: Computer Applications in Bioprocessin. Vol. 70, p. 109 Büssow, K. see Eickhoff, H.: Vol. 77, p. 103 Butler, C. E., Orgill, D. P.: Simultaneous In Vivo Regeneration of Neodermis, Epidermis, and Basement Membrane. Vol. 94, p. 23 Butler, C. E. see Orgill, D. P.: Vol. 93, p. 161 Byun, S. Y. see Choi, J. W.: Vol. 72, p. 63 Cabral, J. M. S. see Fernandes, P.: Vol. 80, p. 115 Cahill, D. J., Nordhoff, E.: Protein Arrays and Their Role in Proteomics. Vol. 83, p. 177 Call, M. K., Tsonis, P. A.: Vertebrate Limb Regeneration. Vol. 93, p. 67 Cantor, C. R. see Jurinke, C.: Vol. 77, p. 57 Cao, N. J. see Gong, C. S.: Vol. 65, p. 207 Cao, N. J. see Tsao, G. T.: Vol. 65, p. 243 Capito, R. M. see Kinner, B.: Vol. 94, p. 91 Carnell, A. J.: Stereoinversions Using Microbial Redox-Reactions. Vol. 63, p. 57 Cash, P.: Proteomics of Bacterial Pathogens. Vol. 83, p. 93 Casqueiro, J. see Martín, J. F.: Vol. 88, p. 91 Cen, P., Xia, L.: Production of Cellulase by Solid-State Fermentation. Vol. 65, p. 69 Chand, S., Mishra, P.: Research and Application of Microbial Enzymes – India’s Contribution. Vol. 85, p. 95 Chang, H. N. see Lee, S. Y.: Vol. 52, p. 27 Chauhan, V. S., Bhardwaj, D.: Current Status of Malaria Vaccine Development. Vol. 84, p. 143 Cheetham, P. S. J.: Combining the Technical Push and the Business Pull for Natural Flavours. Vol. 55, p. 1 Cheetham, P. S. J.: Bioprocesses for the Manufacture of Ingredients for Foods and Cosmetics. Vol. 86, p. 83 Chen, C. see Yang, S.-T.: Vol. 87, p. 61 Chen, Z. see Ho, N. W. Y.: Vol. 65, p. 163 Chenchik, A. see Zhumabayeva, B.: Vol. 86, p. 191 Cho, S. H. see Bruckheimer, E. M.: Vol. 62, p. 75 Cho, G. H. see Choi, J. W.: Vol. 72, p. 63 Choi, J. see Lee, S. Y.: Vol. 71, p. 183 Choi, J. W., Cho, G. H., Byun, S. Y., Kim, D.-I.: Integrated Bioprocessing for Plant Cultures. Vol. 72, p. 63 Christensen, B., Nielsen, J.: Metabolic Network Analysis – A Powerful Tool in Metabolic Engineering. Vol. 66, p. 209 Christians, F. C. see McGall, G. H.: Vol. 77, p. 21 Christmann, M. see Hassfeld, J.: Vol. 97, p. 133 Chu, J. see Zhang, S.: Vol. 87, p. 97

266

Author Index Volumes 51–97

Chu, K. H., Tang, C. Y., Wu, A., Leung, P. S. C.: Seafood Allergy: Lessons from Clinical Symptoms, Immunological Mechanisms and Molecular Biology. Vol. 97, p. 205 Chui, G. see Drmanac, R.: Vol. 77, p. 75 Ciaramella, M. see van der Oost, J.: Vol. 61, p. 87 Colwell, A. S., Longaker, M. T., Lorenz, H. P.: Mammalian Fetal Organ Regeneration. Vol. 93, p. 83 Contreras, B. see Sablon, E.: Vol. 68, p. 21 Conway de Macario, E., Macario, A. J. L.: Molecular Biology of Stress Genes in Methanogens: Potential for Bioreactor Technology. Vol. 81, p. 95 Cordero Otero, R. R. see Hahn-Hägerdal, B.: Vol. 73, p. 53 Cordwell S. J. see Nouwens, A. S.: Vol. 83, p. 117 Cornet, J.-F., Dussap, C. G., Gros, J.-B.: Kinetics and Energetics of Photosynthetic MicroOrganisms in Photobioreactors. Vol. 59, p. 153 da Costa, M. S., Santos, H., Galinski, E. A.: An Overview of the Role and Diversity of Compatible Solutes in Bacteria and Archaea. Vol. 61, p. 117 Cotter, T. G. see McKenna, S. L.: Vol. 62, p. 1 Croteau, R. see McCaskill, D.: Vol. 55, p. 107 Danielsson, B. see Xie, B.: Vol. 64, p. 1 DaPra, E. see Beyeler, W.: Vol. 70, p. 139 Darzynkiewicz, Z., Traganos, F.: Measurement of Apoptosis. Vol. 62, p. 33 Davey, H. M. see Shaw, A. D.: Vol. 66, p. 83 Dean, J. F. D., LaFayette, P. R., Eriksson, K.-E. L., Merkle, S. A.: Forest Tree Biotechnolgy. Vol. 57, p. 1 Debabov, V. G.: The Threonine Story. Vol. 79, p. 113 Debashish, G., Malay, S., Barindra, S., Joydeep, M.: Marine Enzymes. Vol. 96, p. 189 DeFrances M. see Michalopoulos, G. K.: Vol. 93, p. 101 Demain, A. L., Fang, A.: The Natural Functions of Secondary Metabolites. Vol. 69, p. 1 Dhar, N. see Tyagi, A. K.: Vol. 84, p. 211 Diamantis, V. I. see Aivasidis, A.: Vol. 92, p. 49 Diaz, R. see Drmanac, R.: Vol. 77, p. 75 Dochain, D., Perrier, M.: Dynamical Modelling, Analysis, Monitoring and Control Design for Nonlinear Bioprocesses. Vol. 56, p. 147 von Döhren, H.: Biochemistry and General Genetics of Nonribosomal Peptide Synthetases in Fungi. Vol. 88, p. 217 Dolfing, J. see Mogensen, A. S.: Vol. 82, p. 69 Drauz K. see Wöltinger, J.: Vol. 92, p. 289 Driessen, A. J. M. see Evers, M. E.: Vol. 88, p. 111 Drmanac, R., Drmanac, S., Chui, G., Diaz, R., Hou, A., Jin, H., Jin, P., Kwon, S., Lacy, S., Moeur, B., Shafto, J., Swanson, D., Ukrainczyk, T., Xu, C., Little, D.: Sequencing by Hybridization (SBH): Advantages, Achievements, and Opportunities. Vol. 77, p. 75 Drmanac, S. see Drmanac, R.: Vol. 77, p. 75 Du, J. see Gong, C. S.: Vol. 65, p. 207 Du, J. see Tsao, G. T.: Vol. 65, p. 243 Dueser, M. see Raghavarao, K. S. M. S.: Vol. 68, p. 139 Dussap, C. G. see Cornet J.-F.: Vol. 59, p. 153 Dutta, M., Borah, M. M., Dutta, N. N.: Adsorptive Separation of b-Lactam Antibiotics: Technological Perspectives. Vol. 86, p. 255 Dutta, N. N. see Ghosh, A. C.: Vol. 56, p. 111 Dutta, N. N. see Sahoo, G. C.: Vol. 75, p. 209

Author Index Volumes 51–97

267

Dutta, N. N. see Dutta, M.: Vol. 86, p. 255 Dynesen, J. see McIntyre, M.: Vol. 73, p. 103 Eggeling, L., Sahm, H., de Graaf, A. A.: Quantifying and Directing Metabolite Flux: Application to Amino Acid Overproduction. Vol. 54, p. 1 Eggeling, L. see de Graaf, A. A.: Vol. 73, p. 9 Eggink, G., see Kessler, B.: Vol. 71, p. 159 Eggink, G., see van der Walle, G. J. M.: Vol. 71, p. 263 Egli, T. see Wick, L. M.: Vol. 89, p. 1 Ehrlich, H. L. see Rusin, P.: Vol. 52, p. 1 Eickhoff, H., Konthur, Z., Lueking, A., Lehrach, H., Walter, G., Nordhoff, E., Nyarsik, L., Büssow, K.: Protein Array Technology: The Tool to Bridge Genomics and Proteomics. Vol. 77, p. 103 Elias, C. B., Joshi, J. B.: Role of Hydrodynamic Shear on Activity and Structure of Proteins. Vol. 59, p. 47 Eliasson, A. see Gunnarsson, N.: Vol. 88, p. 137 Ellegaard, L. see Angelidaki, I.: Vol. 82, p. 1 Elling, L.: Glycobiotechnology: Enzymes for the Synthesis of Nucleotide Sugars. Vol. 58, p. 89 Enfors, S.-O. see Rozkov, A.: Vol. 89, p. 163 Eriksson, K.-E. L. see Kuhad, R. C.: Vol. 57, p. 45 Eriksson, K.-E. L. see Dean, J. F. D.: Vol. 57, p. 1 Evers, M. E., Trip, H., van den Berg, M. A., Bovenberg, R. A. L., Driessen, A. J. M.: Compartmentalization and Transport in b-Lactam Antibiotics Biosynthesis. Vol. 88, p. 111 Faber, K. see Orru, R. V. A.: Vol. 63, p. 145 Fahnert, B., Lilie, H., Neubauer, P.: Inclusion Bodies: Formation and Utilisation. Vol. 89, p. 93 Fang, A. see Demain, A. L.: Vol. 69, p. 1 Farrace, M. G. see Autuori, F.: Vol. 62, p. 129 Farrell, R. L., Hata, K., Wall, M. B.: Solving Pitch Problems in Pulp and Paper Processes. Vol. 57, p. 197 Fawcett, J. see Verma, P.: Vol. 94, p. 43 Fellenberg, K. see Brazma, A.: Vol. 77, p. 113 Fernandes, P., Prazeres, D. M. F., Cabral, J. M. S.: Membrane-Assisted Extractive Bioconversions. Vol. 80, p. 115 Ferro, A., Gefell, M., Kjelgren, R., Lipson, D. S., Zollinger, N., Jackson, S.: Maintaining Hydraulic Control Using Deep Rooted Tree Systems. Vol. 78, p. 125 Fiechter, A.: Biotechnology in Switzerland and a Glance at Germany. Vol. 69, p. 175 Fiechter, A. see Ochsner, U. A.: Vol. 53, p. 89 Flechas, F. W., Latady, M.: Regulatory Evaluation and Acceptance Issues for Phytotechnology Projects. Vol. 78, p. 171 Foody, B. see Tolan, J. S.: Vol. 65, p. 41 Fréchet, J. M. J. see Xie, S.: Vol. 76, p. 87 Freitag, R., Hórvath, C.: Chromatography in the Downstream Processing of Biotechnological Products. Vol. 53, p. 17 Friehs, K.: Plasmid Copy Number and Plasmid Stability. Vol. 86, p. 47 Furstoss, R. see Orru, R. V. A.: Vol. 63, p. 145 Gadella, T. W. J. see van Munster, E. B.: Vol. 95, p. 143 Le Gal, Y. see Guérard, F.: Vol. 96, p. 127

268

Author Index Volumes 51–97

Galinski, E. A. see da Costa, M. S.: Vol. 61, p. 117 Gàrdonyi, M. see Hahn-Hägerdal, B.: Vol. 73, p. 53 Gatfield, I. L.: Biotechnological Production of Flavour-Active Lactones. Vol. 55, p. 221 Gavala, H. N., Angelidaki, I., Ahring, B. K.: Kinetics and Modeling of Anaerobic Digestion Process. Vol. 81, p. 57 Gavala, H. N. see Skiadas, I. V.: Vol. 82, p. 35 Gefell, M. see Ferro, A.: Vol. 78, p. 125 Gehrke, A. see Brakhage, A. A.: Vol. 88, p. 45 Gemeiner, P. see Stefuca, V.: Vol. 64, p. 69 Gerlach, S. R. see Schügerl, K.: Vol. 60, p. 195 Ghose, T. K., Bisaria, V. S.: Development of Biotechnology in India. Vol. 69, p. 71 Ghose, T. K. see Ghosh, P.: Vol. 85, p. 1 Ghosh, A. C., Mathur, R. K., Dutta, N. N.: Extraction and Purification of Cephalosporin Antibiotics. Vol. 56, p. 111 Ghosh, P., Ghose, T. K.: Bioethanol in India: Recent Past and Emerging Future. Vol. 85, p. 1 Ghosh, P. see Singh, A.: Vol. 51, p. 47 Giesecke, U. see Barber, M. S.: Vol. 88, p. 179 Gilbert, R. J. see Shaw, A. D.: Vol. 66, p. 83 Gill, R. T. see Stephanopoulos, G.: Vol. 73, p. 1 Gomes, J., Menawat, A. S.: Fed-Batch Bioproduction of Spectinomycin. Vol. 59, p. 1 Gong, C. S., Cao, N. J., Du, J., Tsao, G. T.: Ethanol Production from Renewable Resources. Vol. 65, p. 207 Gong, C. S. see Tsao, G. T.: Vol. 65, p. 243 Goodacre, R. see Shaw, A. D.: Vol. 66, p. 83 de Graaf, A. A., Eggeling, L., Sahm, H.: Metabolic Engineering for L-Lysine Production by Corynebacterium glutamicum. Vol. 73, p. 9 de Graaf, A. A. see Eggeling, L.: Vol. 54, p. 1 de Graaf, A. A. see Weuster-Botz, D.: Vol. 54, p. 75 de Graaf, A. A. see Wiechert, W.: Vol. 54, p. 109 Grabley, S., Thiericke, R.: Bioactive Agents from Natural Sources: Trends in Discovery and Application. Vol. 64, p. 101 Gräf, R., Rietdorf, J., Zimmermann, T.: Live Cell Spinning Disk Microscopy. Vol. 95, p. 57 Griengl, H. see Johnson, D. V.: Vol. 63, p. 31 Gros, J.-B. see Larroche, C.: Vol. 55, p. 179 Gros, J.-B. see Cornet, J. F.: Vol. 59, p. 153 Gu, M. B., Mitchell, R. J., Kim, B. C.: Whole-Cell-Based Biosensors for Environmental Biomonitoring and Application. Vol. 87, p. 269 Guenette M. see Tolan, J. S.: Vol. 57, p. 289 Guérard, F., Sellos, D., Le Gal, Y.: Fish and Shellfish Upgrading, Traceability. Vol. 96, p. 127 Gunnarsson, N., Eliasson, A., Nielsen, J.: Control of Fluxes Towards Antibiotics and the Role of Primary Metabolism in Production of Antibiotics. Vol. 88, p. 137 Gupta, M. N. see Roy, I.: Vol. 86, p. 159 Gupta, S. K.: Status of Immunodiagnosis and Immunocontraceptive Vaccines in India. Vol. 85, p. 181 Gutman, A. L., Shapira, M.: Synthetic Applications of Enzymatic Reactions in Organic Solvents. Vol. 52, p. 87

Author Index Volumes 51–97

269

Haagensen, F. see Mogensen, A. S.: Vol. 82, p. 69 Hahn-Hägerdal, B., Wahlbom, C. F., Gárdonyi, M., van Zyl, W. H., Cordero Otero, R. R., Jönsson, L. J.: Metabolic Engineering of Saccharomyces cerevisiae for Xylose Utilization. Vol. 73, p. 53 Haigh, J. R. see Linden, J. C.: Vol. 72, p. 27 Hall, D. O. see Markov, S. A.: Vol. 52, p. 59 Hall, P. see Mosier, N. S.: Vol. 65, p. 23 Hammar, F.: History of Modern Genetics in Germany. Vol. 75, p. 1 Hanai, T., Honda, H.: Application of Knowledge Information Processing Methods to Biochemical Engineering, Biomedical and Bioinformatics Field. Vol. 91, p. 51 Hannenhalli, S., Hubbell, E., Lipshutz, R., Pevzner, P. A.: Combinatorial Algorithms for Design of DNA Arrays. Vol. 77, p. 1 Haralampidis, D., Trojanowska, M., Osbourn, A. E.: Biosynthesis of Triterpenoid Saponins in Plants. Vol. 75, p. 31 Häring, D. see Adam, E.: Vol. 63, p. 73 Harvey, N. L., Kumar, S.: The Role of Caspases in Apoptosis. Vol. 62, p. 107 Hasegawa, S., Shimizu, K.: Noninferior Periodic Operation of Bioreactor Systems. Vol. 51, p. 91 Hassfeld, J., Kalesse, M., Stellfeld, T., Christmann, M.: Asymmetric Total Synthesis of Complex Marine Natural Products. Vol. 97, p. 133 Hata, K. see Farrell, R. L.: Vol. 57, p. 197 Hatton, M. P., Rubin, P. A. D.: Conjunctival Regeneration. Vol. 94, p. 125 Hecker, M.: A Proteomic View of Cell Physiology of Bacillus subtilis – Bringing the Genome Sequence to Life. Vol. 83, p. 57 Hecker, M. see Schweder, T.: Vol. 89, p. 47 van der Heijden, R. see Memelink, J.: Vol. 72, p. 103 Hein, S. see Steinbüchel, A.: Vol. 71, p. 81 Hembach, T. see Ochsner, U. A.: Vol. 53, p. 89 Henzler, H.-J.: Particle Stress in Bioreactor. Vol. 67, p. 35 Herrler, M. see Zhumabayeva, B.: Vol. 86, p. 191 Herrmann, J. see Bruckheimer, E. M.: Vol. 62, p. 75 Hewitt, C. J., Nebe-Von-Caron, G.: The Application of Multi-Parameter Flow Cytometry to Monitor Individual Microbial Cell Physiological State. Vol. 89, p. 197 Hill, D. C., Wrigley, S. K., Nisbet, L. J.: Novel Screen Methodologies for Identification of New Microbial Metabolites with Pharmacological Activity. Vol. 59, p. 73 Hiroto, M. see Inada, Y.: Vol. 52, p. 129 Ho, N. W. Y., Chen, Z., Brainard, A. P., Sedlak, M.: Successful Design and Development of Genetically Engineering Saccharomyces Yeasts for Effective Cofermentation of Glucose and Xylose from Cellulosic Biomass to Fuel Ethanol. Vol. 65, p. 163 Hoch, U. see Adam, W.: Vol. 63, p. 73 Hoff, B. see Schmitt, E. K.: Vol. 88, p. 1 Hoffmann, F., Rinas, U.: Stress Induced by Recombinant Protein Production in Escherichia coli. Vol. 89, p. 73 Hoffmann, F., Rinas, U.: Roles of Heat-Shock Chaperones in the Production of Recombinant Proteins in Escherichia coli. Vol. 89, p. 143 Hofman-Bang, J., Zheng, D., Westermann, P., Ahring, B. K., Raskin, L.: Molecular Ecology of Anaerobic Reactor Systems. Vol. 81, p. 151 Hoheisel, J. see Brazma, A.: Vol. 77, p. 113 Holló, J., Kralovánsky, U. P.: Biotechnology in Hungary. Vol. 69, p. 151

270

Author Index Volumes 51–97

Honda, H., Kobayashi, T.: Industrial Application of Fuzzy Control in Bioprocesses. Vol. 87, p. 151 Honda, H., Liu, C., Kobayashi, T.: Large-Scale Plant Micropropagation. Vol. 72, p. 157 Honda, H. see Hanai, T.: Vol. 91, p. 51 Honda, H., Kobayashi, T.: Large-Scale Micropropagation System of Plant Cells. Vol. 91, p. 105 Hórvath, C. see Freitag, R.: Vol. 53, p. 17 Hou, A. see Drmanac, R.: Vol. 77, p. 75 Houtsmuller, A. B.: Fluorescence Recovery After Photoleaching: Application to Nuclear Proteins. Vol. 95, p. 177 Hubbell, E. see Hannenhalli, S.: Vol. 77, p. 1 Hubbuch, J., Thömmes, J., Kula, M.-R.: Biochemical Engineering Aspects of Expanded Bed Adsorption. Vol. 92, p. 101 Huebner, S. see Mueller, U.: Vol. 79, p. 137 Hummel, W.: New Alcohol Dehydrogenases for the Synthesis of Chiral Compounds. Vol. 58, p. 145 Hüners, M. see Lang, S.: Vol. 97, p. 29 Iijima, S. see Miyake, K.: Vol. 90, p. 89 Iijima, S. see Kamihira, M.: Vol. 91, p. 171 Ikeda, M.: Amino Acid Production Processes. Vol. 79, p. 1 Imamoglu, S.: Simulated Moving Bed Chromatography (SMB) for Application in Bioseparation. Vol. 76, p. 211 Inada, Y., Matsushima, A., Hiroto, M., Nishimura, H., Kodera, Y.: Chemical Modifications of Proteins with Polyethylen Glycols. Vol. 52, p. 129 Irwin, D. C. see Wilson, D. B.: Vol. 65, p. 1 Isermann, H. P. see Bungay, H. R.: Vol. 70, p. 109 Ito, A. see Shinkai, M.: Vol. 91, p. 191 Iwasaki, Y., Yamane, T.: Enzymatic Synthesis of Structured Lipids. Vol. 90, p. 151 Iyer, P. see Lee, Y. Y.: Vol. 65, p. 93 Jackson, S. see Ferro, A.: Vol. 78, p. 125 James, E., Lee, J. M.: The Production of Foreign Proteins from Genetically Modified Plant Cells. Vol. 72, p. 127 Jeffries, T. W., Shi, N.-Q.: Genetic Engineering for Improved Xylose Fementation by Yeasts. Vol. 65, p. 117 Jendrossek, D.: Microbial Degradation of Polyesters. Vol. 71, p. 293 Jenne, M. see Schmalzriedt, S.: Vol. 80, p. 19 Jin, H. see Drmanac, R.: Vol. 77, p. 75 Jin, P. see Drmanac, R.: Vol. 77, p. 75 Johnson, D. V., Griengl, H.: Biocatalytic Applications of Hydroxynitrile. Vol. 63, p. 31 Johnson, E. A., Schroeder, W. A.: Microbial Carotenoids. Vol. 53, p. 119 Johnsurd, S. C.: Biotechnolgy for Solving Slime Problems in the Pulp and Paper Industry. Vol. 57, p. 311 Johri, B. N., Sharma, A., Virdi, J. S.: Rhizobacterial Diversity in India and its Influence on Soil and Plant Health. Vol. 84, p. 49 Jönsson, L. J. see Hahn-Hägerdal, B.: Vol. 73, p. 53 Joshi, J. B. see Elias, C. B.: Vol. 59, p. 47 Joydeep, M. see Debashish, G.: Vol. 96, p. 189 Jurinke, C., van den Boom, D., Cantor, C. R., Köster, H.: The Use of MassARRAY Technology for High Throughput Genotyping. Vol. 77, p. 57

Author Index Volumes 51–97

271

Kaderbhai, N. see Shaw, A. D.: Vol. 66, p. 83 Kalesse, M. see Hassfeld, J.: Vol. 97, p. 133 Kamihira, M., Nishijima, K., Iijima, S.: Transgenic Birds for the Production of Recombinant Proteins. Vol. 91, p. 171 Karanth, N. G. see Krishna, S. H.: Vol. 75, p. 119 Karau, A. see Wöltinger, J.: Vol. 92, p. 289 Karthikeyan, R., Kulakow, P. A.: Soil Plant Microbe Interactions in Phytoremediation. Vol. 78, p. 51 Kataoka, M. see Shimizu, S.: Vol. 58, p. 45 Kataoka, M. see Shimizu, S.: Vol. 63, p. 109 Katzen, R., Tsao, G. T.: A View of the History of Biochemical Engineering. Vol. 70, p. 77 Kawai, F.: Breakdown of Plastics and Polymers by Microorganisms. Vol. 52, p. 151 Kawarasaki, Y. see Nakano, H.: Vol. 90, p. 135 Kell, D. B. see Shaw, A. D.: Vol. 66, p. 83 Kessler, B., Weusthuis, R., Witholt, B., Eggink, G.: Production of Microbial Polyesters: Fermentation and Downstream Processes. Vol. 71, p. 159 Khosla, C. see McDaniel, R.: Vol. 73, p. 31 Khurana, J. P. see Tyagi, A. K.: Vol. 84, p. 91 Kieran, P. M., Malone, D. M., MacLoughlin, P. F.: Effects of Hydrodynamic and Interfacial Forces on Plant Cell Suspension Systems. Vol. 67, p. 139 Kijne, J. W. see Memelink, J.: Vol. 72, p. 103 Kim, B. C. see Gu, M. B.: Vol. 87, p. 269 Kim, D.-I. see Choi, J. W.: Vol. 72, p. 63 Kim, R. see Banks, M. K.: Vol. 78, p. 75 Kim, Y. B., Lenz, R. W.: Polyesters from Microorganisms. Vol. 71, p. 51 Kimura, E.: Metabolic Engineering of Glutamate Production. Vol. 79, p. 37 King, R.: Mathematical Modelling of the Morphology of Streptomyces Species. Vol. 60, p. 95 Kinner, B., Capito, R. M., Spector, M.: Regeneration of Articular Cartilage. Vol. 94, p. 91 Kino-oka, M., Nagatome, H., Taya, M.: Characterization and Application of Plant Hairy Roots Endowed with Photosynthetic Functions. Vol. 72, p. 183 Kino-oka, M., Taya M.: Development of Culture Techniques of Keratinocytes for Skin Graft Production. Vol. 91, p. 135 Kirk, T. K. see Akhtar, M.: Vol. 57, p. 159 Kjelgren, R. see Ferro, A.: Vol. 78, p. 125 Knoll, A., Maier, B., Tscherrig, H., Büchs, J.: The Oxygen Mass Transfer, Carbon Dioxide Inhibition, Heat Removal, and the Energy and Cost Efficiencies of High Pressure Fermentation. Vol. 92, p. 77 Knorre, W. A. see Bocker, H.: Vol. 70, p. 35 Kobayashi, M. see Shimizu, S.: Vol. 58, p. 45 Kobayashi, S., Uyama, H.: In vitro Biosynthesis of Polyesters. Vol. 71, p. 241 Kobayashi, T. see Honda, H.: Vol. 72, p. 157 Kobayashi, T. see Honda, H.: Vol. 87, p. 151 Kobayashi, T. see Honda, H.: Vol. 91, p. 105 Kodera, F. see Inada, Y.: Vol. 52, p. 129 Kohl, T., Schwille, P.: Fluorescence Correlation Spectroscopy with Autofluorescent Proteins. Vol. 95, p. 107 Kolattukudy, P. E.: Polyesters in Higher Plants. Vol. 71, p. 1 König, A. see Riedel, K.: Vol. 75, p. 81 de Koning, G. J. M. see van der Walle, G. A. M.: Vol. 71, p. 263 Konthur, Z. see Eickhoff, H.: Vol. 77, p. 103

272

Author Index Volumes 51–97

Koo, Y.-M. see Lee, S.-M.: Vol. 87, p. 173 Kornprobst, J.-M. see Bourguet-Kondracki, M.-L.: Vol. 97, p. 105 Kossen, N. W. F.: The Morphology of Filamentous Fungi. Vol. 70, p. 1 Köster, H. see Jurinke, C.: Vol. 77, p. 57 Koutinas, A. A. see Webb, C.: Vol. 87, p. 195 Krabben, P., Nielsen, J.: Modeling the Mycelium Morphology of Penicilium Species in Submerged Cultures. Vol. 60, p. 125 Kralovánszky, U. P. see Holló, J.: Vol. 69, p. 151 Krämer, R.: Analysis and Modeling of Substrate Uptake and Product Release by Procaryotic and Eucaryotik Cells. Vol. 54, p. 31 Kretzmer, G.: Influence of Stress on Adherent Cells. Vol. 67, p. 123 Krieger, N. see Mitchell, D. A.: Vol. 68, p. 61 Krishna, S. H., Srinivas, N. D., Raghavarao, K. S. M. S., Karanth, N. G.: Reverse Micellar Extraction for Downstream Processeing of Proteins/Enzymes. Vol. 75, p. 119 Kück, U. see Schmitt, E. K.: Vol. 88, p. 1 Kuhad, R. C., Singh, A., Eriksson, K.-E. L.: Microorganisms and Enzymes Involved in the Degradation of Plant Cell Walls. Vol. 57, p. 45 Kuhad, R. Ch. see Singh, A.: Vol. 51, p. 47 Kula, M.-R. see Hubbuch, J.: Vol. 92, p. 101 Kulakow, P. A. see Karthikeyan, R.: Vol. 78, p. 51 Kulakow, P. A. see Banks, M. K.: Vol. 78, p. 75 Kumagai, H.: Microbial Production of Amino Acids in Japan. Vol. 69, p. 71 Kumar, R. see Mukhopadhyay, A.: Vol. 86, p. 215 Kumar, S. see Harvey, N. L.: Vol. 62, p. 107 Kunze, G. see Riedel, K.: Vol. 75, p. 81 Kwon, S. see Drmanac, R.: Vol. 77, p. 75 Lacy, S. see Drmanac, R.: Vol. 77, p. 75 Ladenstein, R., Antranikian, G.: Proteins from Hyperthermophiles: Stability and Enzamatic Catalysis Close to the Boiling Point of Water. Vol. 61, p. 37 Ladisch, C. M. see Mosier, N. S.: Vol. 65, p. 23 Ladisch, M. R. see Mosier, N. S.: Vol. 65, p. 23 LaFayette, P. R. see Dean, J. F. D.: Vol. 57, p. 1 Lalk, M. see Schweder, T.: Vol. 96, p. 1 Lammers, F., Scheper, T.: Thermal Biosensors in Biotechnology. Vol. 64, p. 35 Lang, S., Hüners, M., Lurtz, V.: Bioprocess Engineering Data on the Cultivation of Marine Prokaryotes and Fungi. Vol. 97, p. 29 Larroche, C., Gros, J.-B.: Special Transformation Processes Using Fungal Spares and Immobilized Cells. Vol. 55, p. 179 Latady, M. see Flechas, F. W.: Vol. 78, p. 171 Lazarus, M. see Adam, W.: Vol. 63, p. 73 Leak, D. J. see van der Werf, M. J.: Vol. 55, p. 147 Lee, J. M. see James, E.: Vol. 72, p. 127 Lee, S.-M., Lin, J., Koo, Y.-M.: Production of Lactic Acid from Paper Sludge by Simultaneous Saccharification and Fermentation. Vol. 87, p. 173 Lee, S. Y., Chang, H. N.: Production of Poly(hydroxyalkanoic Acid). Vol. 52, p. 27 Lee, S. Y., Choi, J.: Production of Microbial Polyester by Fermentation of Recombinant Microorganisms. Vol. 71, p. 183 Lee, Y. Y., Iyer, P., Torget, R. W.: Dilute-Acid Hydrolysis of Lignocellulosic Biomass. Vol. 65, p. 93

Author Index Volumes 51–97

273

Lehrach, H. see Eickhoff, H.: Vol. 77, p. 103 Lenz, R. W. see Kim, Y. B.: Vol. 71, p. 51 Leuchtenberger, W. see Wöltinger, J.: Vol. 92, p. 289 Leung, P. S. C. see Chu, K. H.: Vol. 97, p. 205 Licari, P. see McDaniel, R.: Vol. 73, p. 31 Liebezeit, G.: Aquaculture of “Non-Food Organisms” for Natural Substance Production. Vol. 97, p. 1 Liese, A.: Technical Application of Biological Principles in Asymmetric Catalysis. Vol. 92, p. 197 Lievense, L. C., van’t Riet, K.: Convective Drying of Bacteria II. Factors Influencing Survival. Vol. 51, p. 71 Lilie, H. see Fahnert, B.: Vol. 89, p. 93 Lin, J. see Lee, S.-M.: Vol. 87, p. 173 Linden, J. C., Haigh, J. R., Mirjalili, N., Phisaphalong, M.: Gas Concentration Effects on Secondary Metabolite Production by Plant Cell Cultures. Vol. 72, p. 27 Lindequist, U. see Schweder, T.: Vol. 96, p. 1 Lipshutz, R. see Hannenhalli, S.: Vol. 77, p. 1 Lipson, D. S. see Ferro, A.: Vol. 78, p. 125 Little, D. see Drmanac, R.: Vol. 77, p. 75 Liu, B. see Banks, M. K.: Vol. 78, p. 75 Liu, C. see Honda, H.: Vol. 72, p. 157 Lohray, B. B.: Medical Biotechnology in India. Vol. 85, p. 215 Longaker, M. T. see Colwell, A. S.: Vol. 93, p. 83 Lorenz, H. P. see Colwell, A. S.: Vol. 93, p. 83 Lueking, A. see Eickhoff, H.: Vol. 77, p. 103 Luo, J. see Yang, S.-T.: Vol. 87, p. 61 Lurtz, V. see Lang, S.: Vol. 97, p. 29 Lyberatos, G. see Pind, P. F.: Vol. 82, p. 135 Mac Loughlin, P. F. see Kieran, P. M.: Vol. 67, p. 139 Macario, A. J. L. see Conway de Macario, E.: Vol. 81, p. 95 Madhusudhan, T. see Mukhopadhyay, A.: Vol. 86, p. 215 Maier, B. see Knoll, A.: Vol. 92, p. 77 Malay, S. see Debashish, G.: Vol. 96, p. 189 Malone, D. M. see Kieran, P. M.: Vol. 67, p. 139 Maloney, S. see Müller, R.: Vol. 61, p. 155 Mandenius, C.-F.: Electronic Noses for Bioreactor Monitoring. Vol. 66, p. 65 Markov, S. A., Bazin, M. J., Hall, D. O.: The Potential of Using Cyanobacteria in Photobioreactors for Hydrogen Production. Vol. 52, p. 59 Marteinsson, V. T. see Prieur, D.: Vol. 61, p. 23 Martín, J. F., Ullán, R. V., Casqueiro, J.: Novel Genes Involved in Cephalosporin Biosynthesis: The Three-component Isopenicillin N Epimerase System. Vol. 88, p. 91 Marx, A. see Pfefferle, W.: Vol. 79, p. 59 Mathur, R. K. see Ghosh, A. C.: Vol. 56, p. 111 Matsunaga, T., Takeyama, H., Miyashita, H., Yokouchi, H.: Marine Microalgae. Vol. 96, p. 165 Matsushima, A. see Inada, Y.: Vol. 52, p. 129 Mauch, K. see Schmalzriedt, S.: Vol. 80, p. 19 Mayer Jr., J. E. see Rabkin-Aikawa, E.: Vol. 94, p. 141 Mazumdar-Shaw, K., Suryanarayan, S.: Commercialization of a Novel Fermentation Concept. Vol. 85, p. 29

274

Author Index Volumes 51–97

McCaskill, D., Croteau, R.: Prospects for the Bioengineering of Isoprenoid Biosynthesis. Vol. 55, p. 107 McDaniel, R., Licari, P., Khosla, C.: Process Development and Metabolic Engineering for the Overproduction of Natural and Unnatural Polyketides. Vol. 73, p. 31 McDonell, T. J. see Bruckheimer, E. M.: Vol. 62, p. 75 McGall, G. H., Christians, F. C.: High-Density GeneChip Oligonucleotide Probe Arrays. Vol. 77, p. 21 McGovern, A. see Shaw, A. D.: Vol. 66, p. 83 McGowan, A. J. see McKenna, S. L.: Vol. 62, p. 1 McIntyre, M., Müller, C., Dynesen, J., Nielsen, J.: Metabolic Engineering of the Aspergillus. Vol. 73, p. 103 McIntyre, T.: Phytoremediation of Heavy Metals from Soils. Vol. 78, p. 97 McKenna, S. L., McGowan, A. J., Cotter, T. G.: Molecular Mechanisms of Programmed Cell Death. Vol. 62, p. 1 McLoughlin, A. J.: Controlled Release of Immobilized Cells as a Strategy to Regulate Ecological Competence of Inocula. Vol. 51, p. 1 Memelink, J., Kijne, J. W., van der Heijden, R., Verpoorte, R.: Genetic Modification of Plant Secondary Metabolite Pathways Using Transcriptional Regulators. Vol. 72, p. 103 Menachem, S. B. see Argyropoulos, D. S.: Vol. 57, p. 127 Menawat, A. S. see Gomes J.: Vol. 59, p. 1 Menge, M. see Mukerjee, J.: Vol. 68, p. 1 Merkle, S. A. see Dean, J. F. D.: Vol. 57, p. 1 Mescher, A. L., Neff, A. W.: Regenerative Capacity and the Developing Immune System. Vol. 93, p. 39 Meyer, H. E. see Sickmann, A.: Vol. 83, p. 141 Michalopoulos, G. K., DeFrances M.: Liver Regeneration. Vol. 93, p. 101 Mikos, A. G. see Mistry, A. S.: Vol. 94, p. 1 Minas, W. see Barber, M. S.: Vol. 88, p. 179 Mirjalili, N. see Linden, J. C.: Vol. 72, p. 27 Mishra, P. see Chand, S.: Vol. 85, p. 95 Mistry, A. S., Mikos, A. G.: Tissue Engineering Strategies for Bone Regeneration. Vol. 94, p. 1 Mitchell, D. A., Berovic, M., Krieger, N.: Biochemical Engineering Aspects of Solid State Bioprocessing. Vol. 68, p. 61 Mitchell, R. J. see Gu, M. B.: Vol. 87, p. 269 Miura, K.: Tracking Movement in Cell Biology. Vol. 95, p. 267 Miyake, K., Iijima, S.: Bacterial Capsular Polysaccharide and Sugar Transferases. Vol. 90, p. 89 Miyashita, H. see Matsunaga, T.: Vol. 96, p. 165 Miyawaki, A., Nagai, T., Mizuno, H.: Engineering Fluorescent Proteins. Vol. 95, p. 1 Mizuno, H. see Miyawaki, A.: Vol. 95, p. 1 Möckel, B. see Pfefferle, W.: Vol. 79, p. 59 Moeur, B. see Drmanac, R.: Vol. 77, p. 75 Mogensen, A. S., Dolfing, J., Haagensen, F., Ahring, B. K.: Potential for Anaerobic Conversion of Xenobiotics. Vol. 82, p. 69 Moore, J. C. see Arnold, F. H.: Vol. 58, p. 1 Moracci, M. see van der Oost, J.: Vol. 61, p. 87 Mosier, N. S., Hall, P., Ladisch, C. M., Ladisch, M. R.: Reaction Kinetics, Molecular Action, and Mechanisms of Cellulolytic Proteins. Vol. 65, p. 23 Mreyen, M. see Sickmann, A.: Vol. 83, p. 141 Mueller, U., Huebner, S.: Economic Aspects of Amino Acids Production. Vol. 79, p. 137

Author Index Volumes 51–97

275

Muffler, K., Ulber R.: Downstream Processing in Marine Biotechnology. Vol. 97, p. 63 Mühlemann, H. M., Bungay, H. R.: Research Perspectives for Bioconversion of Scrap Paper. Vol. 65, p. 193 Mukherjee, J., Menge, M.: Progress and Prospects of Ergot Alkaloid Research. Vol. 68, p. 1 Mukhopadhyay, A.: Inclusion Bodies and Purification of Proteins in Biologically Active Forms. Vol. 56, p. 61 Mukhopadhyay, A. see Bhatia, P. K.: Vol. 64, p. 155 Mukhopadhyay, A., Basu, S. K.: Intracellular Delivery of Drugs to Macrophages. Vol. 84, p. 183 Mukhopadhyay, A., Madhusudhan, T., Kumar, R.: Hematopoietic Stem Cells: Clinical Requirements and Developments in Ex-Vivo Culture. Vol. 86, p. 215 Müller, C. see McIntyre, M.: Vol. 73, p. 103 Müller, M., Wolberg, M., Schubert, T.: Enzyme-Catalyzed Regio- and Enantioselective Ketone Reductions. Vol. 92, p. 261 Müller, R., Antranikian, G., Maloney, S., Sharp, R.: Thermophilic Degradation of Environmental Pollutants. Vol. 61, p. 155 Müllner, S.: The Impact of Proteomics on Products and Processes. Vol. 83, p. 1 van Munster, E. B., Gadella, T. W. J.: Fluorescence Lifetime Imaging Microscopy (FLIM), Vol. 95, p. 143 Nagai, T. see Miyawaki, A.: Vol. 95, p. 1 Nagatome, H. see Kino-oka, M.: Vol. 72, p. 183 Nagy, E.: Three-Phase Oxygen Absorption and its Effect on Fermentation. Vol. 75, p. 51 Nakano, H., Kawarasaki, Y., Yamane, T.: Cell-free Protein Synthesis Systems: Increasing their Performance and Applications. Vol. 90, p. 135 Nakashimada, Y. see Nishio, N.: Vol. 90, p. 63 Nath, S.: Molecular Mechanisms of Energy Transduction in Cells: Engineering Applications and Biological Implications. Vol. 85, p. 125 Nebe-Von-Caron, G. see Hewitt, C. J.: Vol. 89, p. 197 Necina, R. see Strancar, A.: Vol. 76, p. 49 Neff, A. W. see Mescher, A. L.: Vol. 93, p. 39 Neubauer, P. see Fahnert, B.: Vol. 89, p. 93 Nielsen, J. see Christensen, B.: Vol. 66, p. 209 Nielsen, J. see Gunnarsson, N.: Vol. 88, p. 137 Nielsen, J. see Krabben, P.: Vol. 60, p. 125 Nielsen, J. see McIntyre, M.: Vol. 73, p. 103 Nisbet, L. J. see Hill, D. C.: Vol. 59, p. 73 Nishijima, K. see Kamihira, M.: Vol. 91, p. 171 Nishimura, H. see Inada, Y.: Vol. 52, p. 123 Nishio, N., Nakashimada, Y.: High Rate Production of Hydrogen/Methane from Various Substrates and Wastes. Vol. 90, p. 63 Nöh, K. see Wiechert, W.: Vol. 92, p. 145 Nordhoff, E. see Cahill, D. J.: Vol. 83, p. 177 Nordhoff, E. see Eickhoff, H.: Vol. 77, p. 103 Nouwens, A. S., Walsh, B. J., Cordwell S. J.: Application of Proteomics to Pseudomonas aeruginosa. Vol. 83, p. 117 Nyarsik, L. see Eickhoff, H.: Vol. 77, p. 103 Ochsner, U. A., Hembach, T., Fiechter, A.: Produktion of Rhamnolipid Biosurfactants. Vol. 53, p. 89

276

Author Index Volumes 51–97

O’Connor, R.: Survival Factors and Apoptosis: Vol. 62, p. 137 Ogawa, J. see Shimizu, S.: Vol. 58, p. 45 Ohshima, T., Sato, M.: Bacterial Sterilization and Intracellular Protein Release by Pulsed Electric Field. Vol. 90, p. 113 Ohta, H.: Biocatalytic Asymmetric Decarboxylation. Vol. 63, p. 1 Oldiges, M., Takors, R.: Applying Metabolic Profiling Techniques for Stimulus-Response Experiments: Chances and Pitfalls. Vol. 92, p. 173 Oliverio, S. see Autuori, F.: Vol. 62, p. 129 van der Oost, J., Ciaramella, M., Moracci, M., Pisani, F. M., Rossi, M., de Vos, W. M.: Molecular Biology of Hyperthermophilic Archaea. Vol. 61, p. 87 Orgill, D. P., Butler, C. E.: Island Grafts: A Model for Studying Skin Regeneration in Isolation from Other Processes. Vol. 93, p. 161 Orgill, D. P. see Butler, C. E.: Vol. 94, p. 23 Orlich, B., Schomäcker, R.: Enzyme Catalysis in Reverse Micelles. Vol. 75, p. 185 Orru, R. V. A., Archelas, A., Furstoss, R., Faber, K.: Epoxide Hydrolases and Their Synthetic Applications. Vol. 63, p. 145 Osbourn, A. E. see Haralampidis, D.: Vol. 75, p. 31 Oude Elferink, S. J. W. H. see Stams, A. J. M.: Vol. 81, p. 31 Padmanaban, G.: Drug Targets in Malaria Parasites. Vol. 84, p. 123 Panda, A. K.: Bioprocessing of Therapeutic Proteins from the Inclusion Bodies of Escherichia coli. Vol. 85, p. 43 Park, E. Y.: Recent Progress in Microbial Cultivation Techniques. Vol. 90, p. 1 Paul, G. C., Thomas, C. R.: Characterisation of Mycelial Morphology Using Image Analysis. Vol. 60, p. 1 Perrier, M. see Dochain, D.: Vol. 56, p. 147 Pevzner, P. A. see Hannenhalli, S.: Vol. 77, p. 1 Pfefferle, W., Möckel, B., Bathe, B., Marx, A.: Biotechnological Manufacture of Lysine. Vol. 79, p. 59 Phisaphalong, M. see Linden, J. C.: Vol. 72, p. 27 Piacentini, G. see Autuori, F.: Vol. 62, p. 129 Pind, P. F., Angelidaki, I., Ahring, B. K., Stamatelatou, K., Lyberatos, G.: Monitoring and Control of Anaerobic Reactors. Vol. 82, p. 135 Piredda, L. see Autuori, F.: Vol. 62, p. 129 Pisani, F. M. see van der Oost, J.: Vol. 61, p. 87 Plattner, H. see Brakhage, A. A.: Vol. 88, p. 45 Podgornik, A. see Strancar, A.: Vol. 76, p. 49 Podgornik, A., Tennikova, T. B.: Chromatographic Reactors Based on Biological Activity. Vol. 76, p. 165 Pohl, M.: Protein Design on Pyruvate Decarboxylase (PDC) by Site-Directed Mutagenesis. Vol. 58, p. 15 Poirier, Y.: Production of Polyesters in Transgenic Plants. Vol. 71, p. 209 Pons, M.-N., Vivier, H.: Beyond Filamentous Species. Vol. 60, p. 61 Pons, M.-N., Vivier, H.: Biomass Quantification by Image Analysis. Vol. 66, p. 133 Prazeres, D. M. F. see Fernandes, P.: Vol. 80, p. 115 Prieur, D., Marteinsson, V. T.: Prokaryotes Living Under Elevated Hydrostatic Pressure. Vol. 61, p. 23 Prior, A. see Wolfgang, J.: Vol. 76, p. 233 Pulz, O., Scheibenbogen, K.: Photobioreactors: Design and Performance with Respect to Light Energy Input. Vol. 59, p. 123

Author Index Volumes 51–97

277

Rabkin-Aikawa, E., Mayer Jr., J. E., Schoen, F. J.: Heart Valve Regeneration. Vol. 94, p. 141 Raghavarao, K. S. M. S., Dueser, M., Todd, P.: Multistage Magnetic and Electrophoretic Extraction of Cells, Particles and Macromolecules. Vol. 68, p. 139 Raghavarao, K. S. M. S. see Krishna, S. H.: Vol. 75, p. 119 Ramanathan, K. see Xie, B.: Vol. 64, p. 1 Raskin, L. see Hofman-Bang, J.: Vol. 81, p. 151 Reichert, A. see Barber, M. S.: Vol. 88, p. 179 Reuss, M. see Schmalzriedt, S.: Vol. 80, p. 19 Riedel, K., Kunze, G., König, A.: Microbial Sensor on a Respiratory Basis for Wastewater Monitoring. Vol. 75, p. 81 van’t Riet, K. see Lievense, L. C.: Vol. 51, p. 71 Rietdorf, J. see Gräf, R.: Vol. 95, p. 57 Rinas, U. see Hoffmann, F.: Vol. 89, p. 73 Rinas, U. see Hoffmann, F.: Vol. 89, p. 143 Roberts, S. M. see Allan, J. V.: Vol. 63, p. 125 Robinson, A. see Brazma, A.: Vol. 77, p. 113 Rock, S. A.: Vegetative Covers for Waste Containment. Vol. 78, p. 157 Roehr, M.: History of Biotechnology in Austria. Vol. 69, p. 125 Rogers, P. L., Shin, H. S., Wang, B.: Biotransformation for L-Ephedrine Production. Vol. 56, p. 33 Rossi, M. see van der Oost, J.: Vol. 61, p. 87 Rowland, J. J. see Shaw, A. D.: Vol. 66, p. 83 Roy, I., Sharma, S., Gupta, M. N.: Smart Biocatalysts: Design and Applications. Vol. 86, p. 159 Roychoudhury, P. K., Srivastava, A., Sahai, V.: Extractive Bioconversion of Lactic Acid. Vol. 53, p. 61 Rozkov, A., Enfors, S.-O.: Analysis and Control of Proteolysis of Recombinant Proteins in Escherichia coli. Vol. 89, p. 163 Rubin, P. A. D. see Hatton, M. P.: Vol. 94, p. 125 Rusin, P., Ehrlich, H. L.: Developments in Microbial Leaching – Mechanisms of Manganese Solubilization. Vol. 52, p. 1 Russell, N. J.: Molecular Adaptations in Psychrophilic Bacteria: Potential for Biotechnological Applications. Vol. 61, p. 1 Sablon, E., Contreras, B., Vandamme, E.: Antimicrobial Peptides of Lactic Acid Bacteria: Mode of Action, Genetics and Biosynthesis. Vol. 68, p. 21 Sahai, V. see Singh, A.: Vol. 51, p. 47 Sahai, V. see Roychoudhury, P. K.: Vol. 53, p. 61 Saha-Möller, C. R. see Adam, W.: Vol. 63, p. 73 Sahm, H. see Eggeling, L.: Vol. 54, p. 1 Sahm, H. see de Graaf, A. A.: Vol. 73, p. 9 Sahoo, G. C., Dutta, N. N.: Perspectives in Liquid Membrane Extraction of Cephalosporin Antibiotics: Vol. 75, p. 209 Saleemuddin, M.: Bioaffinity Based Immobilization of Enzymes. Vol. 64, p. 203 Santos, H. see da Costa, M. S.: Vol. 61, p. 117 Sarkans, U. see Brazma, A.: Vol. 77, p. 113 Sarkiss, M. see Bruckheimer, E. M.: Vol. 62, p. 75 Sato, M. see Ohshima, T.: Vol. 90, p. 113 Sauer, U.: Evolutionary Engineering of Industrially Important Microbial Phenotypes. Vol. 73, p. 129 Scheibenbogen, K. see Pulz, O.: Vol. 59, p. 123

278

Author Index Volumes 51–97

Scheper, T. see Lammers, F.: Vol. 64, p. 35 Schmalzriedt, S., Jenne, M., Mauch, K., Reuss, M.: Integration of Physiology and Fluid Dynamics. Vol. 80, p. 19 Schmidt, J. E. see Skiadas, I. V.: Vol. 82, p. 35 Schmitt, E. K., Hoff, B., Kück, U.: Regulation of Cephalosporin Biosynthesis. Vol. 88, p. 1 Schneider, K. see Beyeler, W.: Vol. 70, p. 139 Schoen, F. J. see Rabkin-Aikawa, E.: Vol. 94, p. 141 Schomäcker, R. see Orlich, B.: Vol. 75, p. 185 Schreier, P.: Enzymes and Flavour Biotechnology. Vol. 55, p. 51 Schreier, P. see Adam, W.: Vol. 63, p. 73 Schroeder, W. A. see Johnson, E. A.: Vol. 53, p. 119 Schubert, T. see Müller, M.: Vol. 92, p. 261 Schubert, W.: Topological Proteomics, Toponomics, MELK-Technology. Vol. 83, p. 189 Schügerl, K.: Extraction of Primary and Secondary Metabolites. Vol. 92, p. 1 Schügerl, K., Gerlach, S. R., Siedenberg, D.: Influence of the Process Parameters on the Morphology and Enzyme Production of Aspergilli. Vol. 60, p. 195 Schügerl, K. see Seidel, G.: Vol. 66, p. 115 Schügerl, K.: Recovery of Proteins and Microorganisms from Cultivation Media by Foam Flotation. Vol. 68, p. 191 Schügerl, K.: Development of Bioreaction Engineering. Vol. 70, p. 41 Schügerl, K. see Tollnick, C.: Vol. 86, p. 1 Schumann, W.: Function and Regulation of Temperature-Inducible Bacterial Proteins on the Cellular Metabolism. Vol. 67, p. 1 Schuster, K. C.: Monitoring the Physiological Status in Bioprocesses on the Cellular Level. Vol. 66, p. 185 Schwab, P. see Banks, M. K.: Vol. 78, p. 75 Schweder, T., Hecker, M.: Monitoring of Stress Responses. Vol. 89, p. 47 Schweder, T., Lindequist, U., Lalk, M.: Screening for New Metabolites from Marine Microorganisms. Vol. 96, p. 1 Schwille, P. see Kohl, T.: Vol. 95, p. 107 Scouroumounis, G. K. see Winterhalter, P.: Vol. 55, p. 73 Scragg, A. H.: The Production of Aromas by Plant Cell Cultures. Vol. 55, p. 239 Sedlak, M. see Ho, N. W. Y.: Vol. 65, p. 163 Seidel, G., Tollnick, C., Beyer, M., Schügerl, K.: On-line and Off-line Monitoring of the Production of Cephalosporin C by Acremonium Chrysogenum. Vol. 66, p. 115 Seidel, G. see Tollnick, C.: Vol. 86, p. 1 Sellos, D. see Guérard, F.: Vol. 96, p. 127 Shafto, J. see Drmanac, R.: Vol. 77, p. 75 Sharma, A. see Johri, B. N.: Vol. 84, p. 49 Sharma, M., Swarup, R.: The Way Ahead – The New Technology in an Old Society. Vol. 84, p. 1 Sharma, S. see Roy, I.: Vol. 86, p. 159 Shamlou, P. A. see Yim, S. S.: Vol. 67, p. 83 Shapira, M. see Gutman, A. L.: Vol. 52, p. 87 Sharp, R. see Müller, R.: Vol. 61, p. 155 Shaw, A. D., Winson, M. K., Woodward, A. M., McGovern, A., Davey, H. M., Kaderbhai, N., Broadhurst, D., Gilbert, R. J., Taylor, J., Timmins, E. M., Alsberg, B. K., Rowland, J. J., Goodacre, R., Kell, D. B.: Rapid Analysis of High-Dimensional Bioprocesses Using Multivariate Spectroscopies and Advanced Chemometrics. Vol. 66, p. 83 Shi, N.-Q. see Jeffries, T. W.: Vol. 65, p. 117

Author Index Volumes 51–97

279

Shimizu, K.: Metabolic Flux Analysis Based on 13C-Labeling Experiments and Integration of the Information with Gene and Protein Expression Patterns. Vol. 91, p. 1 Shimizu, K. see Hasegawa, S.: Vol. 51, p. 91 Shimizu, S., Ogawa, J., Kataoka, M., Kobayashi, M.: Screening of Novel Microbial for the Enzymes Production of Biologically and Chemically Useful Compounds. Vol. 58, p. 45 Shimizu, S., Kataoka, M.: Production of Chiral C3- and C4-Units by Microbial Enzymes. Vol. 63, p. 109 Shin, H. S. see Rogers, P. L.: Vol. 56, p. 33 Shinkai, M., Ito, A.: Functional Magnetic Particles for Medical Application. Vol. 91, p. 191 Sibarita, J.-B.: Deconvolution Microscopy. Vol. 95, p. 201 Sickmann, A., Mreyen, M., Meyer, H. E.: Mass Spectrometry – a Key Technology in Proteome Research. Vol. 83, p. 141 Siebert, P. D. see Zhumabayeva, B.: Vol. 86, p. 191 Siedenberg, D. see Schügerl, K.: Vol. 60, p. 195 Singh, A., Kuhad, R. Ch., Sahai, V., Ghosh, P.: Evaluation of Biomass. Vol. 51, p. 47 Singh, A. see Kuhad, R. C.: Vol. 57, p. 45 Singh, R. P., Al-Rubeai, M.: Apoptosis and Bioprocess Technology. Vol. 62, p. 167 Skiadas, I. V., Gavala, H. N., Schmidt, J. E., Ahring, B. K.: Anaerobic Granular Sludge and Biofilm Reactors. Vol. 82, p. 35 Smith, J. S. see Banks, M. K.: Vol. 78, p. 75 Sohail, M., Southern, E. M.: Oligonucleotide Scanning Arrays: Application to High-Throughput Screening for Effective Antisense Reagents and the Study of Nucleic Acid Interactions. Vol. 77, p. 43 Sonnleitner, B.: New Concepts for Quantitative Bioprocess Research and Development. Vol. 54, p. 155 Sonnleitner, B.: Instrumentation of Biotechnological Processes. Vol. 66, p. 1 Southern, E. M. see Sohail, M.: Vol. 77, p. 43 Spector, M. see Kinner, B.: Vol. 94, p. 91 Spröte, P. see Brakhage, A. A.: Vol. 88, p. 45 Srinivas, N. D. see Krishna, S. H.: Vol. 75, p. 119 Srivastava, A. see Roychoudhury, P. K.: Vol. 53, p. 61 Stafford, D. E., Yanagimachi, K. S., Stephanopoulos, G.: Metabolic Engineering of Indene Bioconversion in Rhodococcus sp. Vol. 73, p. 85 Stamatelatou, K. see Pind, P. F.: Vol. 82, p. 135 Stams, A. J. M., Oude Elferink, S. J. W. H., Westermann, P.: Metabolic Interactions Between Methanogenic Consortia and Anaerobic Respiring Bacteria. Vol. 81, p. 31 Stark, D., von Stockar, U.: In Situ Product Removal (ISPR) in Whole Cell Biotechnology During the Last Twenty Years. Vol. 80, p. 149 Stefuca, V., Gemeiner, P.: Investigation of Catalytic Properties of Immobilized Enzymes and Cells by Flow Microcalorimetry. Vol. 64, p. 69 Steinbüchel, A., Hein, S.: Biochemical and Molecular Basis of Microbial Synthesis of Polyhydroxyalkanoates in Microorganisms. Vol. 71, p. 81 Stellfeld, T. see Hassfeld, J.: Vol. 97, p. 133 Stephanopoulos, G., Gill, R. T.: After a Decade of Progress, an Expanded Role for Metabolic Engineering. Vol. 73, p. 1 Stephanopoulos, G. see Stafford, D. E.: Vol. 73, p. 85 von Stockar, U., van der Wielen, L. A. M.: Back to Basics: Thermodynamics in Biochemical Engineering. Vol. 80, p. 1 von Stockar, U. see Stark, D.: Vol. 80, p. 149 Stocum, D. L.: Stem Cells in CNS and Cardiac Regeneration. Vol. 93, p. 135

280

Author Index Volumes 51–97

Straathof, A. J. J. see Bruggink, A.: Vol. 80, p. 69 Strancar, A., Podgornik, A., Barut, M., Necina, R.: Short Monolithic Columns as Stationary Phases for Biochromatography. Vol. 76, p. 49 Suehara, K., Yano, T.: Bioprocess Monitoring Using Near-Infrared Spectroscopy. Vol. 90, p. 173 Sun, C.-K.: Higher Harmonic Generation Microscopy. Vol. 95, p. 17 Suryanarayan, S. see Mazumdar-Shaw, K.: Vol. 85, p. 29 Suurnäkki, A., Tenkanen, M., Buchert, J., Viikari, L.: Hemicellulases in the Bleaching of Chemical Pulp. Vol. 57, p. 261 Svec, F.: Capillary Electrochromatography: a Rapidly Emerging Separation Method. Vol. 76, p. 1 Svec, F. see Xie, S.: Vol. 76, p. 87 Swanson, D. see Drmanac, R.: Vol. 77, p. 75 Swarup, R. see Sharma, M.: Vol. 84, p. 1 Tabata, H.: Paclitaxel Production by Plant-Cell-Culture Technology. Vol. 87, p. 1 Takeyama, H. see Matsunaga, T.: Vol. 96, p. 165 Takors, R. see Oldiges, M.: Vol. 92, p. 173 Tanaka, T. see Taniguchi, M.: Vol. 90, p. 35 Tang, C. Y. see Chu, K. H.: Vol. 97, p. 205 Tang, Y.-J. see Zhong, J.-J.: Vol. 87, p. 25 Taniguchi, M., Tanaka, T.: Clarification of Interactions Among Microorganisms and Development of Co-culture System for Production of Useful Substances. Vol. 90, p. 35 Taya, M. see Kino-oka, M.: Vol. 72, p. 183 Taya, M. see Kino-oka, M.: Vol. 91, p. 135 Taylor, J. see Shaw, A. D.: Vol. 66, p. 83 Tenkanen, M. see Suurnäkki, A.: Vol. 57, p. 261 Tennikova, T. B. see Podgornik, A.: Vol. 76, p. 165 Thiericke, R. see Grabely, S.: Vol. 64, p. 101 Thomas, C. R. see Paul, G. C.: Vol. 60, p. 1 Thömmes, J.: Fluidized Bed Adsorption as a Primary Recovery Step in Protein Purification. Vol. 58, p. 185 Thömmes, J. see Hubbuch, J.: Vol. 92, p. 101 Timmens, E. M. see Shaw, A. D.: Vol. 66, p. 83 Todd, P. see Raghavarao, K. S. M. S.: Vol. 68, p. 139 Tolan, J. S., Guenette, M.: Using Enzymes in Pulp Bleaching: Mill Applications. Vol. 57, p. 289 Tolan, J. S., Foody, B.: Cellulase from Submerged Fermentation. Vol. 65, p. 41 Tollnick, C. see Seidel, G.: Vol. 66, p. 115 Tollnick, C., Seidel, G., Beyer, M., Schügerl, K.: Investigations of the Production of Cephalosporin C by Acremonium chrysogenum. Vol. 86, p. 1 Torget, R. W. see Lee, Y. Y.: Vol. 65, p. 93 Traganos, F. see Darzynkiewicz, Z.: Vol. 62, p. 33 Trip, H. see Evers, M. E.: Vol. 88, p. 111 Trojanowska, M. see Haralampidis, D.: Vol. 75, p. 31 Tsao, D. T.: Overview of Phytotechnologies. Vol. 78, p. 1 Tsao, G. T., Cao, N. J., Du, J., Gong, C. S.: Production of Multifunctional Organic Acids from Renewable Resources. Vol. 65, p. 243 Tsao, G. T. see Gong, C. S.: Vol. 65, p. 207 Tsao, G. T. see Katzen, R.: Vol. 70, p. 77 Tscherrig, H. see Knoll, A.: Vol. 92, p. 77

Author Index Volumes 51–97

281

Tsonis, P. A. see Call, M. K.: Vol. 93, p. 67 Tüncher, A. see Brakhage, A. A.: Vol. 88, p. 45 Tyagi, A. K., Dhar, N.: Recent Advances in Tuberculosis Research in India. Vol. 84, p. 211 Tyagi, A. K., Khurana, J. P.: Plant Molecular Biology and Biotechnology Research in the Post-Recombinant DNA Era. Vol. 84, p. 91 Ueda, M. see Wazawa, T.: Vol. 95, p. 77 Ukrainczyk, T. see Drmanac, R.: Vol. 77, p. 75 Ulber R. see Muffler, K.: Vol. 97, p. 63 Ullán, R. V. see Martín, J. F.: Vol. 88, p. 91 Uozumi, N.: Large-Scale Production of Hairy Root. Vol. 91, p. 75 Uyama, H. see Kobayashi, S.: Vol. 71, p. 241 VanBogelen, R. A.: Probing the Molecular Physiology of the Microbial Organism, Escherichia coli using Proteomics. Vol. 83, p. 27 Vandamme, E. see Sablon, E.: Vol. 68, p. 21 Vasic-Racki, D. see Wichmann, R.: Vol. 92, p. 225 Verma, P., Fawcett, J.: Spinal Cord Regeneration. Vol. 94, p. 43 Verpoorte, R. see Memelink, J.: Vol. 72, p. 103 Viikari, L. see Suurnäkki, A.: Vol. 57, p. 261 Vilo, J. see Brazma, A.: Vol. 77, p. 113 Vingron, M. see Brazma, A.: Vol. 77, p. 113 Virdi, J. S. see Johri, B. N: Vol. 84, p. 49 Vivier, H. see Pons, M.-N.: Vol. 60, p. 61 Vivier, H. see Pons, M.-N.: Vol. 66, p. 133 Vorgias, C. E. see Antranikian, G.: Vol. 96, p. 219 de Vos, W. M. see van der Oost, J.: Vol. 61, p. 87 Wahlbom, C. F. see Hahn-Hägerdal, B.: Vol. 73, p. 53 Wall, M. B. see Farrell, R. L.: Vol. 57, p. 197 van der Walle, G. A. M., de Koning, G. J. M., Weusthuis, R. A., Eggink, G.: Properties, Modifications and Applications of Biopolyester. Vol. 71, p. 263 Walsh, B. J. see Nouwens, A. S.: Vol. 83, p. 117 Walter, G. see Eickhoff, H.: Vol. 77, p. 103 Wang, B. see Rogers, P. L.: Vol. 56, p. 33 Wang, R. see Webb, C.: Vol. 87, p. 195 Wazawa, T., Ueda, M.: Total Internal Reflection Fluorescence Microscopy in Single Molecule Nanobioscience. Vol. 95, p. 77 Webb, C., Koutinas, A. A., Wang, R.: Developing a Sustainable Bioprocessing Strategy Based on a Generic Feedstock. Vol. 87, p. 195 Weichold, O. see Adam, W.: Vol. 63, p. 73 van der Werf, M. J., de Bont, J. A. M. Leak, D. J.: Opportunities in Microbial Biotransformation of Monoterpenes. Vol. 55, p. 147 Westermann, P. see Hofman-Bang, J.: Vol. 81, p. 151 Westermann, P. see Stams, A. J. M.: Vol. 81, p. 31 Weuster-Botz, D., de Graaf, A. A.: Reaction Engineering Methods to Study Intracellular Metabolite Concentrations. Vol. 54, p. 75 Weuster-Botz, D.: Parallel Reactor Systems for Bioprocess Development. Vol. 92, p. 125 Weusthuis, R. see Kessler, B.: Vol. 71, p. 159 Weusthuis, R. A. see van der Walle, G. J. M.: Vol. 71, p. 263

282

Author Index Volumes 51–97

Wichmann, R., Vasic-Racki, D.: Cofactor Regeneration at the Lab Scale. Vol. 92, p. 225 Wick, L. M., Egli, T.: Molecular Components of Physiological Stress Responses in Escherichia coli. Vol. 89, p. 1 Wiechert, W., de Graaf, A. A.: In Vivo Stationary Flux Analysis by 13C-Labeling Experiments. Vol. 54, p. 109 Wiechert, W., Nöh, K.: From Stationary to Instationary Metabolic Flux Analysis. Vol. 92, p. 145 van der Wielen, L. A. M. see Bruggink, A.: Vol. 80, p. 69 van der Wielen, L. A. M. see von Stockar, U.: Vol. 80, p. 1 Wiesmann, U.: Biological Nitrogen Removal from Wastewater. Vol. 51, p. 113 Williamson, N. M. see Allan, J. V.: Vol. 63, p. 125 Wilson, D. B., Irwin, D. C.: Genetics and Properties of Cellulases. Vol. 65, p. 1 Winson, M. K. see Shaw, A. D.: Vol. 66, p. 83 Winterhalter, P., Skouroumounis, G. K.: Glycoconjugated Aroma Compounds: Occurence, Role and Biotechnological Transformation. Vol. 55, p. 73 Witholt, B. see Kessler, B.: Vol. 71, p. 159 Wolberg, M. see Müller, M.: Vol. 92, p. 261 Wolfgang, J., Prior, A.: Continuous Annular Chromatography. Vol. 76, p. 233 Wöltinger, J., Karau, A., Leuchtenberger, W., Drauz K.: Membrane Reactors at Degussa. Vol. 92, p. 289 Woodley, J. M.: Advances in Enzyme Technology – UK Contributions. Vol. 70, p. 93 Woodward, A. M. see Shaw, A. D.: Vol. 66, p. 83 Wrigley, S. K. see Hill, D. C.: Vol. 59, p. 73 Wu, A. see Chu, K. H.: Vol. 97, p. 205 Xia, L. see Cen, P.: Vol. 65, p. 69 Xie, B., Ramanathan, K., Danielsson, B.: Principles of Enzyme Thermistor Systems: Applications to Biomedical and Other Measurements. Vol. 64, p. 1 Xie, S., Allington, R. W., Fréchet, J. M. J., Svec, F.: Porous Polymer Monoliths: An Alternative to Classical Beads. Vol. 76, p. 87 Xu, C. see Drmanac, R.: Vol. 77, p. 75 Yamane, T. see Iwasaki, Y.: Vol. 90, p. 135 Yamane, T. see Nakano, H.: Vol. 90, p. 89 Yanagimachi, K. S. see Stafford, D. E.: Vol. 73, p. 85 Yang, S.-T., Luo, J., Chen, C.: A Fibrous-Bed Bioreactor for Continuous Production of Monoclonal Antibody by Hybridoma. Vol. 87, p. 61 Yannas, I. V.: Facts and Theories of Induced Organ Regeneration. Vol. 93, p. 1 Yannas, I. V. see Zhang, M.: Vol. 94, p. 67 Yano, T. see Suehara, K.: Vol. 90, p. 173 Yim, S. S., Shamlou, P. A.: The Engineering Effects of Fluids Flow and Freely Suspended Biological Macro-Materials and Macromolecules. Vol. 67, p. 83 Yokouchi, H. see Matsunaga, T.: Vol. 96, p. 165 Zhang, S., Chu, J., Zhuang, Y.: A Multi-Scale Study on Industrial Fermentation Processes and Their Optimization. Vol. 87, p. 97 Zhang, M., Yannas, I. V.: Peripheral Nerve Regeneration. Vol. 94, p. 67 Zheng, D. see Hofman-Bang, J.: Vol. 81, p. 151 Zhong, J.-J.: Biochemical Engineering of the Production of Plant-Specific Secondary Metabolites by Cell Suspension Cultures. Vol. 72, p. 1

Author Index Volumes 51–97

283

Zhong, J.-J., Tang, Y.-J.: Submerged Cultivation of Medicinal Mushrooms for Production of Valuable Bioactive Metabolites. Vol. 87, p. 25 Zhuang, Y. see Zhang, S.: Vol. 87, p. 97 Zhumabayeva, B., Chenchik, A., Siebert, P. D., Herrler, M.: Disease Profiling Arrays: Reverse Format cDNA Arrays Complimentary to Microarrays. Vol. 86, p. 191 Zimmermann, T.: Spectral Imaging and Linear Unmixing in Light Microscopy. Vol. 95, p. 245 Zimmermann, T. see Gräf, R.: Vol. 95, p. 57 Zollinger, N. see Ferro, A.: Vol. 78, p. 125 van Zyl, W. H. see Hahn-Hägerdal, B.: Vol. 73, p. 53

Subject Index

Agarase 196 Alcohol dehydrogenases, extremophiles 244, 247 Algal taxonomy, FAs 57 Alkaline phosphatase, fish 130 Almelysin 196 Alteramide 32 Amidases, thermoactive 248 Amylases 210 Amylases, heat-stable 235 Anchovy, fish meal 111 Angiotensin-converting enzyme (ACE) 141 Antibacterial compounds 41 Antibiotics 35 Anticancer drugs 172 Antifungal compounds 35 Anti-inflammatory agents 38 Antimicrobial drug screening 24 Antioxidants, marine hydrolysates 143 Antitumor compounds, marine 32–34 Antiviral agents 37 Arachidonic acid 171 Attention-deficit disorder (ADHD), PUFA 81 Bacillariophytes 57 Bacteria, fatty acids 54 – PUFA 91 – cell factories 96 Bacterial artificial chromosome 19 Biological screening, marine 28 C-reactive protein 80 Calcitonin, waste hydrolysates 142 Catalases 197 Cathepsins 210 Cellobiose phosphorylase 208 Cellulases 227

Ceramides, Coolia monolis 59 – sponges 67 CGRP, waste hydrolysates 142 CGTase 238 Chemical screening 28 Chitinases 189, 196, 206 Chitin-degrading enzymes 234 Chlorella spp., anti-inflammatory 169 Chloroperoxidase 197 Chlorophytes 169 Clams, lipids 76 Cloning, screening 19 Cnidaria, lipids 69 CO2 fixation, microalgae 182 Coccolithophorids, CO2 fixation 182 Coccoliths 172 Coelenterates, lipids 69 Copepods, lipids 61 Crustaceans, lipids 78 Cryptophytes 170 CTAB detergent 150 Curacin A 34 Cu-Zn superoxide dismutase 197 Cyanobacteria, fatty acids 54 – polysaccharides 167 Cytokines 38 Cytotoxic agents, marine 32 DHA, fatty acids 79, 171 Diatoms 57, 170 Dimethyl sulfide 198 Dinoflagellates 57, 171 DMSP lyase 198 DNA ligase, thermostable 252 DNA microarrays, genomic 23 DNA polymerases 207, 249 DNA sequencing 251 Docosahexaenoic acids (DHA) 49, 79, 171 – fatty acids 79, 171

286 Docosatetraenoic acids (DTA) 171 Dolastatin-10 34 Drug development, genomics, marine 24 Echinoderms, lipids 72 Eicosapentaenoic acid (EPA) 49, 170 Endoglucanase 229 Endothelin 39 Enkephalins 140 Environmental genomics 18 Enzymes, cloning 202, 208 – fish 130 – inhibitors 39 EPA, fatty acids 79, 170 Epolactaene 39 Epoxy-pseudoisoeugenol (2-methylbutyrate) (EPB) 146 Esterases, extremophiles 194, 244, 247 Ethylene 176 Euglenophytes 173 Extremophiles 192, 219 Fatty acids 49 – acetylenic, sponges 66 – biosynthesis, marine algae 55 – non-methylene-interrupted 53 – polyunsaturated 78 – ∆5,9- 62 FISH 18, 23, 26 Fish, genetic traceability 146 Fish oil, PUFA 90, 106, 111 Fish protein hydrolysates 131 Fish silage 131 Fosmids 19 Galactose-1-phosphate uridylyltransferase 210 Gastrin, waste hydrolysates 142 Gene transfer, microalgae 173 Glucoamylases, heat-stable 235 Glucose dehydrogenase 197 Glucose isomerases, extremophiles 244 β-Glucosidase 196 Glucosidases, heat-stable 235 Glutamine synthetase 198 Glycosphingolipids 67 Gorgonians, lipids 69 Growth hormone releasing factors 142 Gymnodinium spp. 172

Subject Index Haloperoxidases 201 Halophiles 227 Halovir 37 Haptophytes 172 Heart disease, PUFA 80 Heterokontophytes 170 High-content screening (HCS) 30 High-throughput screening (HTS) 30 Hormonal regulating-peptides, waste hydrolysates 142 Hydrolases 196, 206 Hydrolysates 128 Hyperthermophilic archaea, proteases 241 Immunopeptides, fish hydrolysates IPNV 208

142

Komodoquinone A 39 Krill lipids 61 LC-PUFA, marine sources 90 LDH, deep-sea fish 130 Leukotriene 83 Ligase chain reaction 252 Ligases 198 Lipases 194 – thermotolerant 244 Lipids 49 Lyases 198 Lycopene k-cyclase 198 Macroalgae, FAs 59 Macrolactins, HIV 37 Malingamide G 56 Maltotriose 237 Mannanase 209 Marine bioprocess engineering 210 Marine genomes, microbial 1, 3 Marinone 35 Massetolides 35 Metabolome analysis 1, 31 Metalloproteinase 196 Methanococcus jannaschii 3, 192, 210 Methoxy FAs 60 Microalgae, gene transfer 173 – mass cultivation 177 – metabolites 165 Molluscs, lipids 73 Monooxygenases 197 Mussels, PUFA 74

Subject Index

287

Neuroactive peptides 140 Nitrite reductase 204 Nitrogenase 205 Nitrous oxide reductase 204 Obionin 39 Octacosapolyenoic FAs 58 One strain-many compounds Opioids 140 Oxidoreductases 197, 204 Oysters, lipids 75

Repeat expansion detection Rhodophytes 168 Riftia pachyptila 17, 194 rRNA genes, cloning 204 RuBisCO 208

41

Patella spp. 75 PCR 249 Pectin-degrading enzymes 232 Pepsins, cold-active fish 131 Pernilase 192 Phomactin D 38 Phospholipids 49 Photobioreactors, microalgae 180 Photoreactor 165 Phytanic acids 64 Phytoplankton, fatty acids 56 Pirellula sp. 17 Plakosides 68 Polyketide synthase (PKS) 90 Porphyra spp. 169 Porphyridium cruentum 57, 169 Prochlorococcus spp. 17 Proteases 189, 241 – gastric, fish 131 Proteolysis, quantification 137 Proteome analysis 1, 22 – drug action 41 Proteorhodopsin, marine bacteria 20 Psoriasis, PUFA 81 Psychrophiles 207, 219, 224 PUFA 49, 79, 170 – biosynthesis 83 – fish 103 – health benefits 79 – heart disease 80 PUFA production, molecular biotechnology 96 Pullulanase 238 Raphidophyceae, FA composition RdRp 208

57

252

Salinamides 38 Salinosporamide A 32 Scallops, lipids 76 Screening, high-throughput 1 Scylla serrata 78 Semiplenamides 39 Shellfish, genetic traceability 146 Shellfish protein hydrolysates 131 Shewanella, PUFA phylogeny 93 Silicatein, silica-synthesizing enzyme Sponges 203, 207 – lipids 62 Squids, lipids 77 Starch-hydrolyzing enzymes 236 Sulfolobus spp. 226, 237 Superoxide dismutases 197, 201 Synechocystis spp. 3

203

Taq polymerase 249 Thermophiles 219, 224, 242 Thermus aquaticus, DNA polymerase 251 Thiotropocin 38 Thraustochytrids 97, 99, 171 Thromboxane 83 Thyrotropin-releasing hormone (TRH) 140 Topoisomerase 172 Transcriptome 1 Transferases 207 Triacylgycerols 51 – biosynthesis 83 Trypsin, fish 130 Tubular reactors, microalgae 180 Tumor necrosis factor 83 Tunicates, lipids 73 VLC-PUFA 58 Xylanolytic enzymes 230 Xylobiose 230 Xylose isomerase 244 Zooplankton, lipids 60