Biotechnological Applications of Extremophilic Microorganisms 9783110424331, 9783110427738

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Natuschka M. Lee (Ed.) Biotechnological Applications of Extremophilic Microorganisms Life in Extreme Environments

Life in Extreme Environments

Series Editor Dirk Wagner

Volume 6

Biotechnological Applications of Extremophilic Microorganisms Edited by Natuschka Lee

Editor Natuschka M. Lee Department of Ecology and Environmental Science Umeå University Linneaus väg 6 901 87 Umeå, Sweden [email protected]

Series Editor Dirk Wagner GFZ German Research Centre for Geosciences, Helmholtz Centre Potsdam Section Geomicrobiology Telegrafenberg 14473 Potsdam, Germany [email protected]

ISBN 978-3-11-042773-8 e-ISBN (PDF) 978-3-11-042433-1 e-ISBN (EPUB) 978-3-11-042436-2 ISSN 2197-9227 Library of Congress Control Number: 2020939096 Bibliographic information published by the Deutsche Nationalbibliothek The Deutsche Nationalbibliothek lists this publication in the Deutsche Nationalbibliografie; detailed bibliographic data are available in the Internet at http://dnb.dnb.de. © 2020 Walter de Gruyter GmbH, Berlin/Boston Cover image: robertcicchetti/iStock/Getty Images Typesetting: Compuscript Ltd., Shannon, Ireland Printing and binding: CPI books GmbH, Leck www.degruyter.com

Preface The Encyclopaedia Britannia defines biotechnology simply as the use of biology to solve problems and make useful products. The term biotechnology was coined in the 1960–1970s when the techniques in cellular and molecular biology were developed. It became obvious that basic research could lead to useful products and processes in the medical and industrial field, such as to produce therapeutic substances or to refine industrial processes by producing useful substances like vitamins and enzymes. In the early days, the most common technique used in biotechnology was genetic engineering. The aim was to introduce genes coding for useful compounds into a “production cell” to enable a large scale production of the desired compound (e.g. insulin). However, with the advancement of basic research in cellular and molecular biology, the development of new powerful instruments, advanced computer technology, and the discovery of extremophilic organisms with astonishing capabilities, a plethora of new applications has emerged in all kinds of scientific fields. It goes without saying that many developments in biotechnology would in fact not have been possible without the extremophiles. The most well-known example is the Taq polymerase, which is used today worldwide in the polymerase chain reaction (PCR). This enzyme was initially isolated from a thermophilic microbe in a hot spring in 1966. Since the late 1970s, the techniques of biotechnology have shifted toward more refined and advanced applications, such as developing preventive measures of diseases, making a process more efficient and sustainable with regard to energy usage or reduced negative impact on the climate, or to even boost up an organism’s survival capabilities. The intention of this volume is to present a survey of the current state of the art and the further potential of biotechnological applications based on different extremophilic or extremotolerant organisms. The contents have been sorted by application field, starting with industrial and medical biotechnology (Chapters 1–5), which is then followed by environmental biotechnology (Chapters 6–9), geobiotechnology (Chapters 10–11), and the emerging new field, astrobiotechnology (Chapters 12–14). In Chapter 1, Kaul and Abouhmad present a survey on how different extremophiles can serve as a source of different novel natural products, while Stennett et al. present in Chapter 2 the amazing pharmacy sources from different extremophiles. In Chapter 3, Karlsson et al. present cutting-edge technology in metabolic engineering of thermophiles, which also includes CRISPR/cas9-based technology. Espina et al., representing an international biotech company, present in Chapter 4 how extremozymes can be screened and used to produce novel substances. In Chapter 5, Kunte et al. demonstrate a new application field, namely how to use ectoine, a compatible solute from halophiles, to support biological material to survive extreme osmotic stress and other stressful conditions, such as freezing and radiation. The applications within environmental biotechnology start with a survey in Chapter 6 by Haruta on how energy is produced and converted in thermophilic https://doi.org/10.1515/9783110424331-202

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photosynthesizing systems and how this can be harnessed based on synthetic ecology. Ferro et al. describe in Chapter 7 how cold-adapted algae can be used to transform wastewater into clean water, air, energy, and useful biosubstances. In Chapter 8, Adeleke et al. describe how microbes can adapt to hazardous substances, such as polycyclic aromatic hydrocarbons, while Þorsteinsdóttir and Vilhelmsson describe in Chapter 9 the potential of bioremediation in cold desert environments. The geobiotechnology section is introduced by Chapter 10, where Stan-Lotter takes us deep down to the subsurface to explore how microbes there can support a safe storage of nuclear waste. Marrero et al. show in Chapter 11 how microbes can be useful for a cost-efficient solution to recover different types of heavy metals from ores and other metal-containing solutions. Last but certainly not least when it comes to amazing novel applications of extremophiles is the new field of astrobiotechnology. Each of the last three chapters deal with a specific group of organisms of great interest for both applications on Earth, as well as in space related research. In Chapter 12, Verseux elaborates how cyanobacteria can be employed on space missions for different purposes, and how lessons learned from this could benefit Earth-based technologies. Vandewalle-Capo et al. reveal in Chapter 13 how even yeast, which was once thought to be just an ordinary mesophile, harbors an impressive potential of tolerance under different extreme conditions. Yeast is therefore not only interesting for different kinds of biotechnological applications, but has also become an interesting model species in astrobiology. The last chapter (14, by Rehamnia et al.) deals with remarkable microscopic animals, the tardigrades – known for their impressive capabilities to survive extreme conditions by morphological adaptations. The biotechnological potential of the tardigrades was actually suggested over 50 years ago, but it is only today, thanks to recent advanced research, that the biotechnological applications of tardigrades have just about started to become truly apparent. More than ever, Louis Pasteur’s statement on the difference between basic research and applied research has shown to be the key solution to all advance in science, in particular in biotechnology: “There does not exist a category of science to which one can give the name applied science. There are science and the applications of science, bound together as the fruit of the tree which bears it.” Umeå, April 8, 2020, Natuschka Lee

Volume published in the series Volume 1 Jens Kallmeyer, Dirk Wagner (Eds.) Micobial Life of the Deep Bisospehere ISBN 978-3-11-030009-3

Volume 2 Corien Bakermans (Ed.) Micobial Evolution under Extreme Conditions ISBN 978-3-11-033506-4

Volume 3 Annette Summers Engel (Ed.) Micobial Life of Cave Systems ISBN 978-3-11-033499-9

Volume 4 Blaire Steven (Ed.) The Biology of Arid Soils ISBN 978-3-11-041998-6

Volume 5 Jens Kallmeyer (Ed.) Life at Vents and Seeps ISBN 978-3-11-049475-4

Volume 8 Étienne Yergeau (Ed.) Advanced Techniques for Studying Microorganisms in Extreme Environments ISBN 978-3-11-052464-2

Contents Preface   v Contributing authors 

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Rajni Hatti-Kaul and Adel Abouhmad Extremophiles: a promising source of novel natural products   1 Tapping the potential of extremophiles for diverse bioactive  compounds   1 1.2 Thermophiles   2 1.3 Cold adapted microbes   5 1.4 Halophiles   9 1.5 Marine extremophiles   13 1.6 Acidophiles   18 1.7 Alkaliphiles   23 1.8 Speeding up the discovery process   26 References   30 1 1.1

Henry L. Stennett, Kavita Tiwari, Sam E. Williams, Paul Curnow and Paul R. Race 2 The extremophilic pharmacy: drug discovery at the limits of life   43 2.1 Introduction   43 2.2 Microorganisms as a source of pharmaceutical leads   43 2.3 Categories of extremophiles   45 2.4 Life on the limit: how environmental conditions drive metabolic innovation   46 2.5 Lomaiviticins   46 2.6 Salinosporamide A   50 2.7 Marinostatin   53 2.8 Abyssomicin C   56 2.9 Macrolactins   61 2.10 Conclusions and future prospects   64 References   64 Eva Nordberg Karlsson, Roya R.R. Sardari, Emanuel Y.C. Ron, Snaedis H. Bjornsdottir, Bjorn T. Adalsteinsson, Olafur H. Fridjonsson and Gudmundur O. Hreggvidsson 3 Metabolic engineering of thermophilic bacteria for production of biotechnologically interesting compounds   73 3.1 Introduction   73 3.2 Increased interest in thermophilic systems in a circular bioeconomy   74

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3.3

Genetic tools and transformation requirements for engineering in thermophiles   77 3.3.1 Competence and transformation   77 3.3.2 Vectors   80 3.3.3 Selection markers   81 3.3.4 Marker recycling   81 3.3.5 CRISPR-Cas9 technology   82 3.4 The importance of metabolic range – anaerobic/aerobic systems for different types of products   83 3.4.1 Thermoanaerobacterium spp. for ethanol or 1,2-propandiol production   85 3.4.2 Geobacillus thermoglucosidasius, a facultative organism engineered for ethanol production   86 3.4.3 Thermotoga spp. and Caldicellulosiruptor bescii, with less developed engineering systems   87 3.4.4 Thermus thermophilus and Rhodothermus marinus – natural pigment producers   88 3.5 Future perspectives   90 References   90 Giannina Espina, Paulina Cáceres-Moreno, Daniela Correa-Llantén, Felipe Sarmiento and Jenny M. Blamey 4 Extremozymes: from discovery to novel bio-products   97 4.1 Abstract   97 4.2 Biocatalysis benefits and barriers   97 4.3 Extremozymes discovery and development functional roadmap   99 4.4 Direct exploration of enzymatic activities   101 4.4.1 Phase 1 – discovery   101 4.4.2 Phase 2 – development   104 4.4.3 Phase 3 – scale-up   109 4.4.4 Phase 4 – production phase   112 4.5 Conclusions   115 References   116 Hans Jörg Kunte, Thomas Schwarz and Erwin A. Galinski 5 The compatible solute ectoine: protection mechanisms, strain development, and industrial production   121 5.1 Introduction   121 5.2 Molecular interaction of ectoine with water and biomolecules and its effect on solution properties   123 5.2.1 Proteins   123 5.2.2 Membranes   125

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5.2.3 Nucleic acids   127 5.2.4 Water relations   129 5.2.5 Solution properties and effects in combination with NaCl   131   5.2.6 Conclusion  132 5.3 Producer strains and fermentation processes   134 5.3.1 Ectoine biosynthesis – genes, enzymes, and regulation   134 5.3.2 Increased ectoine synthesis by improving the expression of ectABC   136 5.3.3 The “bacterial milking” procedure   137 5.3.4 The “leaky” mutant procedure (exploiting a mutant which excretes ectoine into the medium)   138 5.3.5 Mechanisms of ectoine export from H. elongata cells   140 5.3.6 Blocking ectoine degradation – the “super-leaky” mutant   141 5.4 Products   143 References   144 Shin Haruta 6 Thermophilic photosynthesis-based microbial communities – energy production and conversion   153   6.1 Introduction  153 6.2 Thermophilic photosynthetic bacteria   153 6.2.1 Oxygenic photosynthetic bacteria – cyanobacteria–   155 6.2.2 Anoxygenic photosynthetic bacteria   155 6.3 Photosynthesis-based microbial communities in terrestrial hot springs   156 6.3.1 Bacterial and archaeal compositions of hot spring microbial mats   157 6.3.2 Layered structure   157 6.3.3 Productivity   159 6.4 Future directions – microflora engineering   160 References   162 Lorenza Ferro, Fernanda H. B. De Miranda Vasconcelos, Francesco G. Gentili and Christiane Funk 7 Photosynthesis at high latitudes – adaptation of photosynthetic microorganisms to Nordic climates   165 7.1 Introduction   165 7.1.1 Photosynthetic microorganisms: characters of a billion years-old story   165 7.1.2 How, what, where? Growing algae in northern environments   168 7.2 Acclimation strategies of photosynthetic microorganisms living at high latitudes   169

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7.2.1 Light acclimation   169 7.2.2 Cold adaptation   174 7.3 The strange case of Clamydomonas raudensis strain UWO241   178 7.4 MicroBioRefine – biomass production and wastewater ­reclamation in Nordic climate   179 7.4.1 Why wastewater?   181 7.4.2 Why flue gases?   182 7.5 Conclusions and final remarks   183 References   184 Rasheed Adeleke, Maryam Bello-Akinosho, Mphekgo Maila and Natuschka M. Lee 8 Roles of extremophiles in the bioremediation of polycyclic aromatic hydrocarbon contaminated soil environment   197 8.1 Introduction   197 8.2 PAHs in soil environment   198 8.2.1 Direct and indirect impacts of PAH on soil environment   199 8.2.2 Sources of PAH in soil environments   200 8.2.3 Toxicity of PAHs creates extreme environment   201   8.3 Removal of PAHs from the environment  202 8.3.1 Different approaches for PAH removal from soil   202 8.3.2 Bioremediation of PAH-polluted sites   203 8.4 How do extremophiles drive the bioremediation process?   205   8.4.1 Aerobic and anaerobic degradation processes  209 8.4.2 Bioremediation technologies   211 8.5 Future perspectives   212 8.5.1 Can other plant-associated microorganisms – endophytes, plant growth promoting rhizobacteria (PGPR), mycorrhizal helper bacteria, and mycorrhizal fungi – assist extremophiles in PAH degradation?   212 8.6 Summary and Conclusions   216 References   216 Guðný Vala Þorsteinsdóttir and Oddur Vilhelmsson 9 Bioremediative potential of bacteria in cold desert environments  9.1 Bioremediation – general considerations   231   9.2 Hydrocarbon degradation  232 9.2.1 Aerobic degradation   233 9.2.2 Anaerobic degradation   233 9.3 Effects of environmental conditions   234   9.4 Microbial life in cold deserts  235 9.4.1 Bioprospecting cold desert soils for hydrocarbon-degrading microbes   236

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9.5 Concluding remarks  References   238

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Helga Stan-Lotter 10 Subsurface extremophiles and nuclear waste storage   243 10.1 Introduction   243 10.2 Subsurface storage and research sites   246   10.2.1 Deep geological facilities  246 10.2.2 Underground rock laboratories   247 10.2.3 Rock salt deposits   248 10.2.4 Advantages of the Waste Isolation Pilot Plant   249   10.3 Microorganisms and radionuclide storage  250 10.4 Removal of uranium and other radionuclides   252 10.4.1 Adsorption to free or immobilized cells   252 10.4.2 Dissimilatory reduction which decreases solubility   253   10.4.3 Halophilic microorganisms and radionuclides  255 10.5 Nuclear transmutation   256 References   257 Jeannette Marrero, Orquidea Coto and Axel Schippers 11 Metal bioleaching: fundamentals and geobiotechnical application of aerobic and anaerobic acidophiles   261 11.1 Introduction   261 11.2 Acidophiles: habitats and adaptation to low pH   262 11.3 Biomining and bioleaching: concepts and mechanisms   264 11.3.1 Sulfide and oxide ores   264 11.3.2 Fundamentals of bioleaching and biooxidation mechanisms   266   11.4 Geobiotechnical application of acidophiles: biomining examples  269 11.4.1 Irrigation-type processes   270 11.4.2 Commercial applications of irrigation-type bioleaching processes: copper sulfide ore bioprocesses   270 11.4.3 Stirred tank-type processes for sulfide minerals   271 11.4.4 Bioleaching of oxide ores   273 References   281 Cyprien Verseux 12 Cyanobacterium-based technologies in space and on Earth  12.1 Introduction   289

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12.2 Cyanobacterium-based life support systems for space exploration   289 12.3 Transferring cyanobacterium-based life-support systems to Earth   291

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12.4 Contributions of cyanobacterium-based processes to carbon fixation   293 12.5 Cyanobacteria as a source of energy   294   12.6 Cyanobacteria for food production  296 12.7 Cyanobacteria against reverse desertification   297 12.8 Other biotechnological applications of cyanobacteria   298 12.9 Engineering cyanobacteria   299 12.10 Conclusion   302 Acknowledgements   302 References   303 Marine Vandewalle-Capo, Eric Capo, Baraa Rehamnia, Merlin Sheldrake and Natuschka M. Lee   313 13 The biotechnological potential of yeast under extreme conditions   313 13.1 Introduction   313 13.1.1 Overview of the fungal kingdom   313 13.1.2 Overview of the yeast biology   314 13.1.3 Cultivation of yeasts   316 13.1.4 Microbial ecology under extreme conditions   316   13.1.5 Global diversity of extremophilic yeast  318 13.2 Yeast diversity in extreme environments   318 13.2.1 Cold environments   318 13.2.2 Hot environments   321   13.2.3 Acidic environments  322 13.2.4 Alkaline environments   322 13.2.5 Saline environments   323 13.2.6 Desiccated environments   324   13.2.7 Deep-sea environments  324 13.2.8 Terrestrial dark environments   325 13.2.9 Environments with radiation   326 13.2.10 Polluted environments   326 13.3 Biotechnological applications of extremophilic yeast   327 13.3.1 Food biotechnology   327 13.3.2 Environmental biotechnology   331 13.3.3 Biofuel   332 13.3.4 Biosynthesis   334 13.3.5 Agriculture and aquaculture   335 13.3.6 Biomedical and pharmaceutical applications   336 13.3.7 Insect biotechnology   338 13.3.8 Astrobiotechnology   339 13.4 Summary   339 References   340

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Baraa Rehamnia, Renaud de Cherisey, Olivier Braissant, and Natuschka Lee  357 Biotechnological potential of tardigrades  Historical overview of the discovery of tardigrades and their cryptobiotic  357 abilities   358 14.2 A brief review of what kind of animals tardigrades are   362 14.3 The characteristics of tolerance in tardigrades   363 14.3.1 Anhydrobiosis   364 14.3.2 Cryobiosis     365 14.3.3 Osmobiosis  366 14.3.4 Anoxybiosis   366 14.4 Biotechnological potential of tardigrades  14.4.1 Preservation molecules and processes in anhydrobiotic  368 organisms   370 14.4.2 Medical biotechnology   373 14.4.3 Environmental biotechnology     375 14.4.4 Tardigrades in space research     14.5 Summary 377  378 References  14 14.1

Index 

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Contributing authors Adel Abouhmad Biotechnology, Center for Chemistry & Chemical Engineering Lund University Naturvetarvägen 14 Box 124 SE-221 00 Lund, Sweden [email protected] Bjorn T. Adalsteinsson Matis ohf Vinlandsleid 12 113 Reykjavik, Iceland [email protected] Rasheed Adeleke Agricultural Research Council Institute of Soil, Climate and Water (ARC-ISCW) 600 Belvedere Street, Pretoria Pretoria 0001, South Africa AND Unit for Environmental Science and Management, North-West University Potchefstroom 2520, South Africa AND Lehrstuhl für Mikrobiologie Technische Universität München Emil Ramann Strasse 4 85354 Freising, Germany [email protected] Maryam Bello-Akinosho Agricultural Research Council Institute of Soil, Climate and Water (ARC-ISCW) 600 Belvedere Street, Pretoria Pretoria 0001, South Africa AND Department of Microbiology and Plant Pathology University of Pretoria, Lynnwood Road Hatfield Pretoria 0002, South Africa [email protected]

Snaedis H Bjornsdottir Department of Biology, University of Iceland, Sturlugata 7, Askja 102 Reykjavik, Iceland [email protected] Jenny M. Blamey Swissaustral USA LLC 111 Riverbend Rd. Office #271 Athens Georgia 30602, USA AND Fundación Científica y Cultural Biociencia Research and Development Jose Domingo Cañas 2280 Ñuñoa 7750132 Santiago, Chile [email protected] Olivier Braissant Department of Biomedical Engineering University of Basel, Switzerland 14 Gwerbestrasse 4123 Allschwil [email protected] Paulina Cáceres-Moreno Swissaustral USA 111 Riverbend Rd. Office #271 Athens GA 30602, USA AND Fundación Científica y Cultural Biociencia Development and Production Jose Domingo Cañas 2280 Ñuñoa 7750132 Santiago, Chile [email protected]

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 Contributing authors

Eric Capo Department of Chemistry Umeå University 901 87 Umeå, Sweden [email protected] Daniela Correa-llantén Swissaustral USA 111 Riverbend Rd. Office #271 Athens GA 30602, USA AND Fundación Científica y Cultural Biociencia Research and Development Jose Domingo Cañas 2280 Ñuñoa 7750132 Santiago, Chile [email protected] Orquidea Coto Federal Institute for Geosciences and Raw Materials Geozentrum Hannover Stilleweg 2 30655 Hannover, Germany [email protected] Paul Curnow School of Biochemistry Biomedical Sciences Building University of Bristol Bristol, BS8 1TD, United Kingdom AND BrisSynBio Synthetic Biology Research Centre Life Sciences Building Tyndall Avenue Bristol, BS8 1TQ, United Kingdom [email protected] Renaud de Cherisey Department of Ecology and Environmental Science Umeå University 6 Linneaus väg 901 87 Umeå, Sweden AND Université de Poitiers 12 Rue du Cuvier 8600 Poitiers, France [email protected]

Giannina Espina Swissaustral USA, 111 Riverbend Rd. Office #271 Athens GA 30602, USA AND Fundación Científica y Cultural Biociencia Research and Development Jose Domingo Cañas 2280 Ñuñoa 7750132 Santiago, Chile [email protected] Lorenza Ferro Department of Chemistry Umeå University 901 87 Umeå, Sweden [email protected] Olafur H. Fridjonsson Matis ohf, Vinlandsleid 12 113 Reykjavik, Iceland [email protected] Christiane Funk Department of Chemistry Umeå University 901 87 Umeå, Sweden [email protected] Erwin Galinski Institute of Microbiology and Biotechnology Rheinische Friedrich-Wilhelms-Universität Meckenheimer Allee 168 53115 Bonn, Germany [email protected] Francesco Gentili Department of Forest Biomaterials and Technology Swedish University of Agricultural Sciences 90183 Umeå, Sweden [email protected] Shin Haruta Department of Biological Sciences Tokyo Metropolitan University 1-1 Minami-Osawa, Hachioji Tokyo 192-0397, Japan [email protected]

Contributing authors 

Rajni Hatti-Kaul Biotechnology, Center for Chemistry & Chemical Engineering Lund University Naturvetarvägen 14 Box 124 SE-221 00 Lund, Sweden [email protected] Gudmundur O Hreggvidsson Matis ohf, Vinlandsleid 12 113 Reykjavik, Iceland AND Department of Biology, University of Iceland Askya 102 Reykjavik, Iceland [email protected] Eva Nordberg Karlsson Biotechnology, Department of Chemistry Lund University PO Box 124 22100 Lund, Sweden [email protected]

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Mphekgo Maila Agricultural Research Council Institute of Soil, Climate and Water (ARC-ISCW) 600 Belvedere Street, Pretoria Pretoria 0001, South Africa [email protected] Jeannette Marrero Federal Institute for Geosciences and Raw Materials Geozentrum Hannover Stilleweg 2 30655 Hannover, Germany [email protected] Fernanda H. B. De Miranda Vasconcelos Department of Wildlife Fish and Environmental Studies Swedish University of Agricultural Sciences, 90183 Umeå, Sweden [email protected]

Hans-Jörg Kunte Federal Institute for Materials Research and Testing (BAM) Division FB 4.1 Biodeterioration and Reference Organisms Unter den Eichen 87 12205 Berlin, Germany [email protected]

Paul R. Race School of Biochemistry Biomedical Sciences Building University of Bristol Bristol, BS8 1TD, United Kingdom AND BrisSynBio Synthetic Biology Research Centre Life Sciences Building Tyndall Avenue Bristol, BS8 1TQ, United Kingdom [email protected]

Natuschka M. Lee Department of Ecology and Environmental Science Umeå University Linneaus väg 6 (KBC) 901 87 Umeå, Sweden [email protected] AND Lehrstuhl für Mikrobiologie, Technische Universität München Emil Ramann Strasse 4 85354 Freising, Germany

Baraa Rehamnia Department of Ecology and Environmental Science Umeå University 6 Linneaus väg 901 87 Umeå, Sweden AND Department of Microbial Biotechnology Université Frères Mentouri Constantine, Algeria [email protected]

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Emanuel Y.C. Ron Biotechnology, Department of Chemistry Lund University PO Box 124 22100 Lund, Sweden [email protected] Roya R.R. Sardari Biotechnology, Department of Chemistry Lund University PO Box 124 22100 Lund, Sweden [email protected] Felipe Sarmiento Swissaustral USA LLC 111 Riverbend Rd. Office #271 Athens Georgia 30602, USA [email protected] Axel Schippers Federal Institute for Geosciences and Raw Materials Geozentrum Hannover Stilleweg 2 30655 Hannover, Germany [email protected] Merlin Sheldrake 20 Willow Road London, NW3 1TJ, United Kingdom [email protected] Helga Stan-Lotter University of Salzburg Department of Biosciences Hellbrunnerstr. 34 5020 Salzburg, Austria [email protected]

Henry L. Stennett School of Biochemistry Biomedical Sciences Building University of Bristol BS8 1TD, United Kingdom AND BrisSynBio Synthetic Biology Research Centre Life Sciences Building Tyndall Avenue Bristol, BS8 1TQ, United Kingdom [email protected] Thomas Schwarz bitop AG Carlo-Schmid-Allee 5, 44263 Dortmund, Germany [email protected] Kavita Tiwari School of Biochemistry Biomedical Sciences Building University of Bristol Bristol, BS8 1TD, United Kingdom [email protected] Guðný Vala Þorsteinsdóttir Icelandic Institute of Natural History – 4 Akureyri Branch Borgir vid Nordurslod 600 Akureyri, Iceland [email protected] Marine Vandewalle-Capo Department of Molecular Biology, Umeå University 90187 Umeå, Sweden [email protected] Cyprien Verseux Laboratory of Applied Space Microbiology ZARM – Center of Applied Space Technology and Microgravity Universität Bremen Am Fallturm 2 28359 Bremen, Germany [email protected]

Contributing authors 

Oddur Vilhelmsson Faculty of Natural Resource Sciences University of Akureyri 2 Borgir vid Nordurslod 600 Akureyri, Iceland AND School of Biological Sciences University of Reading Earley Reading RG6 6AS, United Kingdom [email protected]

Sam E. Williams School of Biochemistry Biomedical Sciences Building University of Bristol Bristol, BS8 1TD, United Kingdom [email protected]

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Rajni Hatti-Kaul and Adel Abouhmad

1 Extremophiles: a promising source of novel natural products 1.1 Tapping the potential of extremophiles for diverse bioactive compounds Extremophiles, the microorganisms inhabiting different extreme environments characterized by high or low temperature, high or low pH, high salt concentration, high pressure, high radiation, etc. or combinations thereof, have developed unique strategies for adapting and thriving in such environments. Studies on extremophiles have been pursued with great interest to determine the mechanisms of adaptation and also as an important source of useful products including enzymes, polymers, compatible solutes, etc. During the last 20–30 years, attention has been directed toward search for novel bioactive compounds produced by extremophiles, which most likely are playing a role in controlling microbial population in the respective ecological niches and are also promising candidates for applications in foods and healthcare. Secondary metabolites produced by microbes have long been a major source of natural products for drug development. Toward the end of the last century, however, high rates of rediscovery of bioactive products from nature prompted a shift to high-throughput screening programs based on molecular targets and combinatorial chemistry, which has unfortunately not led to major discoveries of novel products, and the trend is now to build focused libraries around the chemical scaffolds of natural products. An urgent need for new molecules that could potentially replace the present-day antibiotics, which are becoming ineffective due to the resistance developed by the pathogens, has served as an important driver for the increasing efforts on mining the unexplored ecological niches for bioactive compounds. The importance of extremophiles, the “non-mesophilic” microorganisms from diverse environments, as a unique source of novel antibacterial, antifungal, and antitumor molecules cannot be overlooked, as evidenced by the increasing number of publications, including reviews and books, especially during the past decade [1–4] and also the number of drugs already developed or under development (primarily from marine sources) [3, 5]. The traditional route of discovering novel products from extremophiles by isolation of microbes and screening for the desired bioactivity has its limitations. Besides missing a major fraction of the unculturable organisms, cultivation of extremophiles under laboratory conditions can be slow and cumbersome due to their extremely slow growth and limited quantities of the active ­molecules produced for further evaluation. Moreover, the need for extreme environmental conditions required for their growth may require special design or material of the equipment suitable for cultivation. These challenges are now being addressed by https://doi.org/10.1515/9783110424331-001

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 1 Extremophiles: a promising source of novel natural products

the possibility to sequence whole genomes of organisms and metagenomes in an environmental sample as well as bioinformatics and analytical technologies. The aim of this chapter is to provide a glimpse of the diversity of bioactive molecules produced by extremophilic microorganisms in different environments and also the impact of the latest technological advances on speeding up the discovery process. Environments with extremes of temperature, pH, and salinity are represented in this chapter. The marine habitats are dealt with separately, as they are a combination of high salinity with extreme temperatures and variable acidity and pressure and most importantly have been a source of a larger number of unique metabolites with novel bioactivity than that obtained from terrestrial microorganisms [3].

1.2 Thermophiles Thermophiles, the microbes inhabiting hot environments, are the group that caught the initial interest of the research community and triggered the exploration of the ­fascinating area of extremophiles. Thermophilic (growing at temperatures of 50–79°C) and hyperthermophilic (growing at 80°C and above) microbes have been isolated from diverse high-temperature environments around the world like hot springs, volcanoes, deep oil wells, deep sea hydrothermal vents, compost heaps, etc. Eubacteria and Archaea are the common microorganisms found, while fungal and algal species are limited in number in such environments. Thermophiles differ from other microorganisms in having novel structures, such as thermostable enzymes, polysaccharides with repeating branched oligosaccharide units, lipids with isoprenoid chains (with 15, 20, 25, or 40 carbons rather than straight chain) and with ether linked glycerol or polyols, modified polar glyco- and phospholipids, isoprenoid quinones, polyamines and modified nucleosides that play a role in stabilizing DNA, and osmolytes (e.g. mannosylglycerate) for protection of biological macromolecules and cells [1, 6–8]. The primary application of thermophiles in industrial biotechnology has been as an important source of enzymes that could be used in reactions and processes requiring high temperatures, e.g. for DNA amplification in polymerase chain reaction, biomass pretreatment for production of biofuels and chemicals, and biocatalysis for chemical transformations for pharmaceutical and fine chemical industries [9–11]. Production of a number of selective pharmaceutically active compounds or building blocks has indeed been achieved using thermostable enzymes from several hyperthermophilic bacteria and archaea, which is either very difficult or not possible using chemical catalysis. Carotenoid pigments are ubiquitous in extremophiles and are of interest as food colorants, dietary supplements, and also for pharmaceuticals. In thermophiles, their major function is likely related to membrane stabilization at high temperatures, besides scavenging reactive oxygen species for cellular protection in many non-phototrophic bacteria [12]. A large fraction of the carotenoids reported from

1.2 Thermophiles 

 3

thermophilic bacteria are in the form of glycoside esters such as thermozeaxanthins, zeaxanthin mono- and diglucoside esters and thermocryptoxanthin in Thermus ther­ mophilus [13, 14], zeaxanthin monoglucoside, thermozeaxanthin and thermobiszeaxanthin in Thermus filiformis [12], keto-myxocoxanthin glucoside fatty acid ester (containing iso fatty acids) in the filamentous phototrophic bacterium Roseiflexus castenholzii [15], and a series of carotenoid esters in red pigmented moderate thermophile Meiothermus ruber [16], thermophilic green sulfur bacterium Chlorobium tepid­ ium [17], and thermophilic halophilic Rhodococcus marinus [18]. Genome sequence analysis of T. thermophilus has revealed that the genes for the enzymes catalyzing the initial steps of carotenoid precursor biosynthesis are scattered around the chromosome, while the genes for the terminal steps are on a plasmid [19], in contrast to T. filiformis, in which all the genes involved in carotenoid biosynthesis are located in the same genomic region constituting a cluster [20]. It was further shown that the carotenoid profile in T. filiformis was influenced as a result of heat shock response, favoring the synthesis of thermozeaxanthins and thermobiszeaxanthins [20]. Concerning the bioactive molecules, since the first report on isolation of Thermorubin (1) an orange red pigment from Thermoactinomyces antibioticus showing antibacterial activity [21], a variety of compounds including alkaloids, peptides (generally termed as bacteriocins), cyclic peptides, polyketides, siderophores, etc. with antibacterial, antifungal, and/or antitumor activities have been reported from thermophilic bacteria (primarily actinomycetes), fungi, and hyperthermophilic archaea, as shown in Tab. 1.1 (see Fig. 1.1 for some representative structures 1–11). Fuscachelins [7], the novel nonribosomal peptide siderophores that chelate ferric ions, were the first secondary metabolites to be identified from thermophiles by genome mining of the moderate thermophile Thermobifida fusca, which led to the identification of the gene cluster for their biosynthesis [30]. The presence of unique sulfur-containing bioactive compounds from thermophilic microbes has been reported. These include BS-1 [4], a trisulfide with cytotoxic activity against several cancer cell

Thermozymocidin (2)

BS-1 (4) Talathermophilin A (9) Dihydrogranaticin (3)

1,2,4,5-Tetrathiane (5)

N-propionyl anthranilic acid (11)

Thermorubrin (1)

Pyochelin (6)

Thermolide A (10)

Malbranpyrrole (8) Fuscachelin-A (7)

Fig. 1.1: Chemical structures of several bioactive compounds isolated from (hyper-)thermophilic microorganisms. See Tab. 1.2 for details.

Bioactivity

Anoxybacillus kamchetkensis Anoxybacillus flavithermus Talaromyces thermophiles YMI3-4

Immunostimulatory effect on carp

Potent nematocidal activity

Herbicidal activity

Thermolides (10)

N-propionyl anthranilic acid (11)

Laceyella sacchari

Talaromyces thermophiles YMI-3

Weak nematocidal activity

Talathermophilins A (9) and B Cyclic dipeptides

Malbranchea sulfurea

Cytotoxic activity against breast and liver cancer cell lines

Malbranpyrrole (8)

Fuscachelins A (7)-C

Pyochelin (6)

[36]

[35]

[33, 34]

[32]

[31]

[30]

[29]

[26, 27] [28]

Japan Hydrothermal system near Obock, Djibouti Italian volcano Yellowstone National Park, USA University of Wisconsin, Madison, USA, 1969 Soil of fumaroles in Sinchong River Hot Springs Zone, Pingtung County, Taiwan Tengchong hot spring, Yunan, China Xi´an hot spring in Shaanxi, China Tengchong hot spring, Yunan, China Green algae, Toyama Prefecture, Japan

[22–24]

[25]

Soil samples in Italy and Canada

Mycelia sterilia, Myriococcum albomyces

[21]

Reference

Fresh horse-swine mixed manure, Berlin, Germany

Soil sample in Pavia, Italy

Location

Thermoactinomyces antibioticus

Thermophile

Streptomyces thermoviolaceus subsp. pigens var WR-141 (ATCC 19283) Bacillus stearothermophilus UK563 Hyperthermophilic archaea Thermococcus tadjuricus T. acidaminovorans Iron binding, potent antifungal activity against Candida Pseudomonas akbaalia albicans, C. glabrata, and Aspergillus fumigatus Chelates ferric ions Thermobifida fusca ATCC 27730

Potent antimicrobial activity against Gram-positive bacteria, low activity against Gram-negative; inhibits protein synthesis, aldose reductase inhibitor Thermozymocidin or Antifungal activity; immunosuppressant; potent Myriocin 2 (2) inhibitor of serine palmitoyltransferase, catalyzing the step in sphingosine synthesis; inhibits proliferation of an interleukin-2-dependent mouse cytotoxic cell line Dihydrogranaticin (3) Antimicrobial activity against Bacillus cereus, inhibits viruses, mycobacteria, and both Gram-positive and -negative eubacteria BS-1 (4) Antitumor activity Cyclic polysulfides (5) Fungicidal activity

Thermorubin (1)

Bioactive compound

Tab. 1.1: Examples of the bioactive compounds isolated from thermophilic microorganisms.

4   1 Extremophiles: a promising source of novel natural products

1.3 Cold adapted microbes 

 5

lines isolated from a strain of thermophilic Bacillus stearothermophilus UK563 strain [26, 27], a compound whose chemical synthesis was patented in 1942 [37], and cyclic polysulfides [5] with fungicidal activity isolated from hyperthermophilic archaea of sulfur-metabolizing strains of the genus Thermococcus [28] (Tab. 1.1). Besides, there are cases of novel metabolites whose bioactivity has not yet been demonstrated, e.g. sibyllimycine, an azaindolizidine metabolite from Thermoactinomycetes sp. from a hot spring at Lake Tanganyika in Cape Banza, Africa [38].

1.3 Cold adapted microbes As several regions of the Earth are exposed to low temperatures over a significant period of time and the fact the polar regions of Antarctic and Arctic and deep seas occupy quite a large area of the planet, the abundance and highly diverse population of cold adapted organisms with strategies to cope with the changing environment is to be expected. Research on cold adapted microorganisms, psychrophiles (with optimum growth temperature below 15°C and maximum around 20°C), and psychrotrophs (opt- and max growth temperatures above 15°C and 20°C, respectively) over the years has shown the presence of cold active enzymes, carotenoids, cryoprotective substances, etc. The usefulness of cold active enzymes is ascribed to their higher catalytic efficiency at low temperatures that would potentially shorten reaction time and prevent undesirable transformations especially in processes containing sensitive components and moreover can be easily inactivated by a moderate increase in temperature [39, 40]. Microorganisms and also multicellular organisms from polar regions, deep sea sediments, alpine region, and other cold environments have attracted interest as a source of natural bioproducts, resulting in the discovery of a vast number of novel natural compounds with biological activities like antibacterial, antifungal, antivirus, antitumor, etc. between 2001 and 2016 [41, 42]. Some examples of the bioactive products produced by eubacteria and fungi are provided in Tab. 1.2. See Fig. 1.2 for structures 12–27. Among the bacterial isolates from polar soils in Antarctica and permafrost soil samples in the Arctic, the most promising sources of antibacterial and antifungal molecules have been Actinobacteria, predominantly psychrotolerant Streptomyces sp. [41, 44, 54–56], but also Arthrobacter, Micromonospora, Nocardioides, Rhodococcus, etc. [57, 58]. Bacterial isolates other than Actinobacteria displaying antimicrobial activity include Bradyrhizobium, Pseudomonas, Enterococcus, Methylobacterium species, and others [41]. Many of the bioactive molecules identified are proteinaceous in nature. Shekh et al. [59] have detected a potent antifungal molecule against the multidrug resistant pathogen Candida albicans in a strain of Enterococcus faecium isolated from an Antarctic penguin rookery, and the activity was ascribed to a 40-amino acid protein of class II bacteriocin. Screening of antimicrobial producers among 8000 cold adapted bacterial isolates (predominantly psychrophilic) from soil samples in Isla de

Rugulosin (22) and skyrin (23)

Flexirubin (21)

Violacein 6 (20)

Chetracins (19)

Griseofulvin (17) Geomycin A-C (18)

Communesins G (16), H

Psychrophilin D Cycloaspeptides A (15), D

Psychrophilin B (14), C

Psychrophilin A

Azalomycin B, Nigericin (13) Non-polyenic macrolide

Streptomyces griseus NTK97 Antimicrobial activity against Gram-positive bacteria Streptomyces no. 8 Antibacterial and antifungal activity against Candida utilis and phytopathogens Penicillium reibeum

Penicillium rivulum IBT24420 Moderate activity against the P388 leukemia cell line Pencillium algidum Moderate antiplasmodial activity P. reibeum P. algidum Cytotoxic P. rivulum (IBT24420) Antifungal activity P. griseofulvum Geomycin B shows antifungal activity against Geomyces sp. A. fumigatus Geomycin C has antibacterial activity against Grampositive S. aureus and Gram-negative E. coli Cytotoxic activity against different cancer cell lines Oidiodendron truncatum GW3-13 Antimycobacterial activity Janthinobacterium sp. Ant5-2 Antimycobacterial activity Flavobacterium sp. Ant342 Antibacterial activity; higher activity against Penicillium Gram-positive pathogens chrysogenum Skyrin shows receptor-selective glucagon IWW1053 antagonism in rat and human hepatocytes

Antifungal and cytotoxic activity

Moderate inhibitor for Gram-positive bacteria

Frigocyclinone (12)

Psychrophile

Bioactivity

Bioactive compound

Tab. 1.2: Bioactive compounds from cold adapted organisms.

[46]

[45]

[44]

[43]

Reference

Soil sample under lichens near Chinese Antarctic Station, Antarctica Benthic mat sample from a fresh water lake, East Antarctica Benthic mat sample from a fresh water lake, East Antarctica Benthic mat sample from a saline Highway Lake in Vestfold Hills, Antarctica

[53]

[52]

[52]

[51]

Soil sample in Greenland [47] Wyoming, USA [45, 47] Greenland IBT Culture Collection, BioCentrum-DTU, [46] Technical University of Denmark [48, 49] Soil sample in Foldes Peninsula and [50] King George island, Antarctica

Soil sample under a redcurrant bush in the tundra region of Wyoming, USA

Soil sample from Terra Nova Bay, Edmunson Point, Antarctica Soil sample from Victoria Land, Antarctica

Location

6   1 Extremophiles: a promising source of novel natural products

1.3 Cold adapted microbes 

Frigocyclinone (12)

 7

Nigericin (13)

Psychrophilin B (14)

Cycloaspeptide A (15)

Communesin G (16)

Violacein (20) Griseofulvin (17)

Geomycin C (18) Chetracin B (19)

Flexirubin (21)

Rugulosin (22)

Skyrin (23)

Usimine A (24) Stereocalpin (25)

Oidioperazine B (26)

Thiopeptin (27)

Fig. 1.2: Chemical structures of selected bioactive compounds isolated from cold adapted organisms. See also Tab. 1.2.

8 

 1 Extremophiles: a promising source of novel natural products

los Estados Reservation (South Atlantic Argentina) led to the selection of psychrotolerant isolates phylogenetically related to Serratia proteamaculans and one psychrophilic Pseudomonas sp. with a wide spectrum of activity against Gram-positive and -negative bacteria [60]. The produced antimicrobial compounds were suspected to be microcin-like (micro-bacteriocin, 3–5 kDa) compounds; a novel potent peptide Serraticin A was purified from a Serratia isolate grown at 8°C, which inhibited bacterial growth at a minimum inhibitory concentration (MIC) of 0.01–0.6 µg/mL depending on the bacteria and medium [61]. Lichens recovered from the polar regions have also attracted attention as source of bioactive compounds, especially considering that they harbor a variety of bacterial communities providing different functions [62]. Earlier studies with the Antarctic lichen Stereocaulon alpinum have shown the presence of Usimines A (24)-C and usinic acid [63], with inhibitory activity against the therapeutically targeted tyrosine phosphatase 1B [64], and then a cyclic depsipeptide stereocalpin A (25) that also showed weak inhibition of tyrosine phosphatase and additionally marginal cytotoxicity against some human solid tumor cell lines [65] (Fig. 1.2). More recently, screening of the microbiota associated with lichens recovered from Arctic and Antarctic by Kim and coworkers [66] have revealed different classes of bacteria, majority of them related to Sphingomonas but also species belonging to other genera, some of which have shown broad range of antibacterial activity against both Gram-positive and -negative pathogens as well as antioxidant activity. Psychrophilic bacteria have even been shown to produce antimicrobial silver nanoparticles, as shown by incubating the culture supernatants of Psychrobacter sp., Aeromonas salmonicida, Pseudomonas veronii, and Yersinia kristensenii isolated from Antarctica, with AgNO3 [67]. The nanoparticles produced at 4°C were in general more active and stable than those produced at 30°C. Among the fungal isolates with antimicrobial activity present in polar soils, Penicillium and Aspergillus species are most prevalent. As seen in Tab. 1.2, several psychrotolerant Penicillium species (P. reibeum, P. rivulum, and P. algidum) isolated from different continents were found to produce psychrophilins (A, B (14), C, D), novel cyclic peptides possessing a nitro group instead of an amino group. On the other hand, P. griseofulvum isolates from separate locations produced griseofulvin (17), a known antifungal compound. In addition, the fungal isolates produced even other bioactive metabolites [45–49]; e.g. P. reibeum and P. algidum produced pentapeptides cycloaspeptide A (15) and D, while P. rivulum produced cytotoxic alkaloids communesins G (16) and H. An array of novel as well as known bioactive molecules have been found in a few other fungal isolates from Antarctic soil samples (Tab. 1.2, Fig. 1.2); e.g. Geomyces sp. produced asterric acid derivatives (geomycins) A–C (geomycin C, 18) [50], while Oidiodendron truncatum GW3-13 produced two new epipolythiodioxopiperazines (chetracins B (19) and C) and five new diketopiperazines (chetracin D and oidioperazines A–D (oidioperazine B, 26)) [51]. Screening of Antarctic mosses as a potential source of novel endophytic fungi and products has led to the isolation of

1.4 Halophiles 

 9

Mortierella alpina from the moss Schistidium antarctici, and its extracts revealed the presence of high amounts of polyunsaturated fatty acids as well as a strong antibacterial activity that was attributed to pyrrolopyrazine alkaloids found in the fungal extract [68]. Reports on microorganisms inhabiting fresh water lakes and ponds in the Polar Regions have been few and localized to Antarctic benthic mats accumulated over thousands of years, undisturbed and without higher metazoans. Members of Cyanobacteria such as Pseudophormidium and Nostoc species are overrepresented in these microbial communities, along with a limited number of eubacterial species and fungal species (predominantly Penicillium species) [41]. Antibacterial and antifungal activities have been detected among several members of these communities. A new cyclic thiazolyl peptide class of antibacterial compounds related to siomycin, cyclothiazomycin, or thiopeptin (27), with potent activity against Gram-positive bacteria including methicillin-resistant bacteria, has been isolated from two strains of Arthrobacter agilis [69], while bioactive pigments violacein 6 (J-PVP) (20) and flexirubin (F-YOP) (21) were isolated from the other psychrophilic eubacteria Janthino­ bacterium sp. Ant5-2 and Flavobacterium sp. Ant342, respectively [52] (Tab. 1.2, Fig. 1.2). Both pigments exhibit antimycobacterial activity against both avirulent and virulent Mycobacterium strains. J-PVP had ~15-fold lower MIC value than that previously reported for violacein pigment from Chromobacterium violaceum. A gene cluster encoding the biosynthesis of polyene moiety of flexirubin in the eubacterium Chitin­ ophaga pinesis, also belonging to the flavobacterium group, has been identified and the enzymes involved in initiating the synthesis of polyene have been characterized [70]. During screening of fungal isolates in the benthic mats, antimicrobial activity was observed in some cold-tolerant species of Penicillium, Aspergillus, Beauvaria, and Cladosporium, and two bioactive bis-anthraquinones, rugolosin (22) and skyrin (23), were identified as products of Penicillium chrysogenum [53] (Tab. 1.2, Fig. 1.2).

1.4 Halophiles Microorganisms inhabiting the salt-rich environments are grouped depending on the range of salt concentration needed for their optimum growth – halotolerant (do not require elevated salt concentration for growth but can grow under saline conditions), moderate halophiles (3–15% w/v), and extreme halophiles (15% w/v up to saturation). The natural hypersaline habitats include salt lakes, salterns, coastal dunes, saline deserts, etc. The largest habitat is the seawater that harbors many halophilic microbial species. The high salinity of several of these environments is combined with extreme temperature and/or pH. To counter the high salt concentration in the environment, halophilic bacteria accumulate a high concentration of compatible solutes, while halophilic archaea accumulate high concentration of salts intracellularly. The compatible solutes, e.g. ectoine and hydroxyectoine, are regarded to have potential

10 

 1 Extremophiles: a promising source of novel natural products

as stabilizers of biomolecules and as stress-protective agents. Their production is achieved by growing the bacteria at high salt concentration, followed by recovering the solutes from the cells by lowering the salinity of the medium [71]. Halophilic bacteria and archaea also constitute an interesting source of enzymes that function in high-salt environments or low water activity, exopolysaccharides, carotenoids, retinal proteins (e.g. rhodopsin), and also polyhydroxyalkanoates, the biodegradable polyesters [72–75]. The main advantage of using extremely halophilic bacteria and archaea is the possibility to grow them under aseptic conditions with low risk of cross-contamination due to the high concentration of salt. Salt lakes and ponds are often colored due to the presence of pigmented ­microorganisms such as halophilic green algae Dunaliella salina rich in β-carotene, haloarchaea producing mainly C50 carotenoids including bacterioruberin and its ­derivatives, and halophilic bacteria, e.g. Salinibacter ruber that produces salinixanthin, a C40 acyl glycoside carotenoid [76, 77]. Other carotenoids found in minor amounts in haloarchaea are lycopersene, cis- and trans-phytofluene, and cisand trans-phytoene. Some species can also produce high amounts of β-carotene, canthanxanthin, and trans-astaxantin. The carotenoids seem to provide protective effect to the cells against damage by the visible and ultraviolet (UV) light, and also reinforcement of the cell membrane [77]. A study comparing the in vitro scavenging capacity of carotenoid pigments against radical and non-radical species showed that the carotenoid extracts of haloarchaea, including bacterioruberin and derivatives, displayed higher antioxidant activity as compared to the zeaxanthins from thermophiles [20]. This was attributed to the presence of acyclic carotenoids with large numbers of both conjugated double bonds and hydroxyl groups in the major carotenoids of the halophiles. Both halophilic Archaea and bacteria produce bioactive peptides, microhalocins and halocins (3–5 to 35 kDa) [78, 79]. Some halocins, especially the ones from Archaea, act on a broad range of microorganisms, while some act on closely related species [78–81]. Halocin sensitivity appears to be more common between strains belonging to different genera irrespective of their location in the same or different saline environments [79, 82]. So far, only a few microcins and halocins have been characterized [78, 83–86]. Their mechanism of action is not fully clear but is probably related to affecting permeability of the cell membrane and resulting in disturbance of ion flux across the membrane [87]. Tab. 1.3 lists several other natural products isolated from halophilic microorganisms, structures of some of which (e.g. 28–37) are shown in Fig. 1.3. Products of peptidic origin other than microcins and halocins include the cyclic peptides (dike­ topiperazines), depsipeptides, and lasso peptides. Cyclo(L-Pro-L-Val) (28) produced by the extremely halophilic archaeon Haloterrigena hispanica isolated from Fuente de Piedra saline lake, Spain, was shown to activate N-acyl homoserine lactone (AHL) bioreporters, indicating that Archaea may have the ability to interact with AHL-­producing bacteria within mixed communities [98]. On the other hand, ­ Haloterrigena sp.

Bioactivity

Moderate cytotoxicity against A-549 cells and weak cytotoxicity against HL-60 cells with IC50 values of 3.0 and 27 μM UV protectant

Erythronolides H Significant antibacterial activity, moderate cytotoxicity (36), I (Congeners of Erythromycin A, Erythromycin C) Streptomonomicin (37) Antimicrobial activity against Gram-positive bacteria, especially Bacillus anthracis

Indole-3-ethenamide (34) Euhalothece-362 (35)

Cyclo(L-Pro-L-Phe) (28), Inhibit bacterial quorum sensing cyclo(L-Pro-L-Leu), cyclo(L-Pro-L-isoLeu) Miuraenamides (29) Inhibition of NADH (nicotinamide adenine dinucleotide, reduced form) oxidase, IC50 (half maximal inhibitory concentration) values of 50 µM Potent inhibition of growth of phytopathogenic oomycete, Phytophthora capsici (MIC 0.4 µg/mL), moderate inhibition against fungi and yeast (6.3–12.5 µg/mL) N-3(-methylbutanoyl)- Cytotoxic activity against murine leukemia P388 cells with IC50 value of 3 µg/mL 3-nitrotyramine (30) Variecolorquinones (31) Moderate to weak cytotoxicity against murine leukemia P388 cells, human promyelocytic leukemia HL-60, and some carcinoma cell lines Variecolorins (32) Weak radical scavenging activity by Variecolorins A-K Bacillamide A (33)-C Algicide

Bioactive compound

Tab. 1.3: Bioactive compounds from halophiles. Location

Reference

Great Salt Plains, Oklahoma, USA Jilantai salt field, Inner Mongolia, China

Streptomonospora alba US Department of Agriculture [97] YIM90003 Agricultural Research Service Culture Collection

[95]

[94]

[92] [93]

[91]

[90]

 11

[96]

Hypersaline microbial mat on San Salvador, Bahamas Aspergillus sclerotorium Putian salt field sediment in Fujian province, China Halophilic unicellular Gypsum crust on the bottom cyanobacterium of a hypersaline saltern Euhalothece sp. LK-1 pond in Eilat, Israel Halophilic actinomycete Dried salt lake in Xingjiang Actinopolyspora sp. province, China YIM90600

Bacillus endophyticus

Halophilic facultatively anaerobic eubacterium Halotolerant fungus Aspergillus variecolor

Hypersaline cyanobacterial [88] mat in the inland desert wadi in South Eastern Oman Moderately halophilic Soil sample near Arai-Hama [89] Paraliomyxa miuraensis beach in Kanagawa, Japan SMH-27-4

Marinobacter sp.

Halophile

1.4 Halophiles 

12 

 1 Extremophiles: a promising source of novel natural products

Cyclo(L-Pro-L-Phe) (28)

Miuraenamide A (29)

N-3-(methylbutanoyl)-3nitrotyramine (30)

Variecolorin A (32)

Variecolorquinone A (31)

Bacillamide A (33)

Euhalothece-362 (35) Indole-3-ethenamide (34) Erythronolide H (36)

Streptomonomicin (37)

Fig. 1.3: Chemical structures of bioactive compounds isolated from halophiles. See also Tab. 1.3.

isolated from a hypersaline cyanobacterial mat, in the inland desert wadi in South Eastern Oman, along with other extremely halophilic and moderately thermophilic bacterial strains identified as Marinobacter and Halomonas, were found to produce antibacterial and quorum sensing inhibitory compounds that can potentially be used as antifouling agents [88]. A family of novel depsipeptides, miuraenamides A–F, β-methoxyacrylate-type antibiotics known to target the electron transfer system of the mitochondrial respiratory chain, was produced by Paraliomyxa miuraensis SMH-27-4 isolated in Japan [89, 99]. Only miuraenamides A (29)–C exhibited potent and selective inhibition against a phytopathogen Phytophthora capsici, suggesting

1.5 Marine extremophiles 

 13

the importance of the conformation of the β-methoxyacrylate group and polyketide moiety of the molecules [99]. Miuraenamide A (29) exerted only moderate antifungal activity [89]. Lasso peptides represent a class of ribosomally synthesized and posttranslationally modified peptides with diverse biological activities, including antimicrobial, enzyme inhibitory, and receptor antagonistic activities [100]. Characterization of an antibiotic lasso peptide, streptomonomicin (37), isolated from a halophilic actinomycete Streptomonospora alba showed the presence of 52% hydrophobic residues and a Se-Asp isopeptide linkage [97]. Some other bioactive compounds isolated from different halophilic bacteria or fungi include indole- and quinone (variecolorquinones)-type (e.g. 31) metabolites, isoechilin-type alkaloids (variecolorins) (e.g. 32), indole (bacillamides (e.g. 33), indole-3-ethenamide (34)) and nitrotyramine (N-3(-methylbutanoyl)-3-nitrotyramine) derivatives (30), macrolide type antibiotics (erythromycin C), and mycosporene-like amino acid (euhalothece-362 (35)), highlighting the biological and structural diversity of halophiles and their metabolites (Tab. 1.3, Fig. 1.3). There are also examples of microorganisms with interesting activity profiles but the representative molecules are not yet known. For example, potent antibacterial and cytotoxic activities were recently reported in fungal strains Aspergillus versicolor, A. terreus, A. flavus, Penicillium purpurogenum, and Eurotium amstelodami isolated from Rann of Kutch, a massive salt desert in the western state of Gujarat, India [101]. In another recent report, the anti-methicillin resistant Staphylococcus aureus potential of halophilic bacteria belonging to several genera, including Bacillus, Halobacillus, Halomonas, Marinobacter, Pseudomonas, Oceanobacillus, Cellulomonas, etc. isolated from Great Salt Plain of Oklahoma, USA, was demonstrated [102].

1.5 Marine extremophiles Marine environments are recognized as the most promising ecosystems having greatest potential for providing access to unique bioactive molecules. Covering more than 70% of the Earth’s surface and with salinity of at least 3%, a span of temperature from near freezing in polar regions and deep seas to extremely high (over 400°C) in deep sea hydrothermal vents, and also variable acidity and pressure, the diversity of marine microorganisms is potentially enormous. The deep sea hydrothermal vents (>200 m) are low in diversity with respect to the macro-organisms but host an enormous microbial diversity. Microorganisms in the hydrothermal vents have been of interest as sources of thermostable enzymes, unusual exopolysaccharides, and lipids. There are a few reports on novel secondary metabolites from vent microbes, e.g. chroman derivatives, ammonificins A (38) and B from Thermovibrio ammonifi­ cans [103], and amphiphilic peptide siderophores, loihichelins A (39)–F from a Halo­ monas sp. [104] (Fig. 1.4).

14 

 1 Extremophiles: a promising source of novel natural products

Diazepinomicin (47)

Loihichelin A (39) Abyssomycin C (46)

g-Indomycinone (45)

Ammonificin A (38)

Spongouridine (42)

Cytarabin (40)

Vidarabine (41)

Halaven (Eribulin mesylate (43)

Salinosporamide (50) Streptokordin (49)

Glyciapyrroles (48)

Trabectedin (44)

Lobophorin H (52) Carboxamycin (51)

Macrolactin A (53)

Mixirin A (54) Curacin A (55) Penicillipyrone A (60) Cochliomycin D (62)

Meleagrin D (59)

Dolastatin 10 (56)

Bryostatin 1 (63) Scytonemin (57)

Plinabulin (58)

Rakicidin D (61)

Fig. 1.4: Chemical structures of several bioactive compounds isolated from marine microorganisms. More information is provided in Tab. 1.4.

During the past two decades, several screening programs aiming at mining of marine habitats, particularly the cold waters of deep seas and polar regions, have resulted in a wealth of novel natural products ranging from simple linear peptides to complex polycyclic polyethers with antibacterial, antifungal, antiviral, antimalarial, antiparasitic, antitumor, and cytotoxic activities from bacteria, fungi, sponges, bryozoans, tunicates, molluscs, and soft corals. Experience so far has shown a significantly higher success rate of drug discovery as compared to that from terrestrial microorganisms. Marine sponges have been ranked at the top with respect to the discovery of novel bioactive molecules. There already exist a handful of approved marine derived drugs, and several more drugs are in different phases of clinical and preclinical trials [105–107]. The Food and Drug Administration–approved drugs cytarabine (40) (Cytostar-U, anticancer) and vidarabine (41) (Vira-A, antiviral) have their origin in the molecules (spongothymidine and spongouridine (42), respectively) isolated from the sponge Tethya crypta in the Carribean Sea [105], and eribulin mesylate (43) (E7389, anticancer) is derived from halochondrin B (a polyether macrolide) isolated from a sponge Axinella sp. [108] (Fig. 1.4). Latruncula sponges inhabiting the cold-water regions of the Southern Hemisphere are a rich source of discorhabdin-type pyrroloiminoquinone alkaloids with strong anticancer activity; however, these compounds have not ­proceeded to

4680 m deep sediment core

Marine sediment sample from 289 m depth, Japan, deposited as DSM15899

Deep sea marine ascidian Didemnum proliferum Kott., Shishijima Island, Japan; Sponge Aplysina aerophoba collected from the Mediterranean sea Marine sediment sample in Alaska

3000 m in the Ayu Trough

Marine sediment, Bahamas

Streptomyces sp.

Verriucosispora sp. AB-18-032

Micromonospora sp. DPJ12; RV115

Streptomyces strain KORDI-3238

Salinispora sp. CNB-440 Streptomyces sp.

3814 m marine sediment, Canary basin Streptomyces sp. South China deep-sea sediment sample Gram positive marine 980 m deep sediment core from bacterium, Bacillus sp. the North Pacific

Streptomyces sp. NPS008187

Location

Microbial isolate

Glyciapyrrole A: IC50 of 180 µM against colorectal adenocarcinoma HT-29 and melanoma B16-F10 human cancer cell line Modest cytotoxicity and broad spectrum antibiotic activity

Streptokordin (49) (methylpyridine derivative) Salinosporamide A* (50) Potent activity against solid tumor and hematologic malignancies, proteasome inhibitor Carboxamycin (51) Inhibitory activity against Gram-positive bacteria and against several tumor cell lines, phosphodiesterase activity lobophorins H (52) and I lobophorins H showed potent activity against Bacillus subtilis Macrolactins A (53)–F Selective antibacterial activity, significant inhibitory effect on mammalian herpes simplex viruses and human melanoma cancer cells

Glyciapyrroles A (48)–C (pyrrolosesquiterpenes)

Diazepinomicin (47)

Pluramycin class of antibiotics

Indomycinone (45) (pluramycin metabolite) Abyssomycin C (46) Potent inhibitor of p-aminobenzoic acid biosynthesis and, hence, folic acid biosynthesis; potent activity against Gram-positive bacteria including the antibiotic resistant Staphylococcus aureus Antibacterial, antioxidative, anti-inflammatory and antitumor activity, broad spectrum in vitro cytotoxicity

Activity

Bioactive product

Tab. 1.4: Examples of some novel metabolites produced by marine microorganisms.

[125]

[124]

[123]

[122]

[121]

[120]

[118, 119]

[117]

[116]

Reference

1.5 Marine extremophiles   15

[132] [133] [134]

Weak cytotoxicity against A-549 cell line Inhibitory effects on human cancer cell lines

Scytonemin* (57) Plinabulin (NPI-2358)* (58) synthetic analogue of Halimide Meroterpenoids Austalide S-U Alkaloids Meleagrin D (59), E Breviane spiroditerpenoids Sesquiterpene γ-pyrones Penicillipyrones A (60) and B Cyclopeptide Rakicidin D (61) Resorcylic acid lactones, Cochliomycins D (62)−F

Waldo lake, Oregon

Cultured from marine algae Halimeda lacrimosa collected in the Bahamas Xisha Island, China

Marine sediment in Samut Sakhon province, Thailand. Southern Chinese Sea

* Compounds that are developed as drugs or undergoing clinical development

Streptomyces sp. MWW064 Cochliobolus lunatus M351

Aspergillus aureolatus HDN14-107 Penicillium sp. F23-2 5080 m ocean sediment sample, China Penicillium sp. 5115 m deep sediment, East Pacific Ocean Penicillium sp. F011 Korean marine sediment

Serine/threonine kinase inhibitor, anti-inflammatory and antiproliferative activities Inhibits tubulin polymerization through binding near colcichin binding site, destabilizes tumor vasculor system, induces direct apoptotic effect on tumor cells Antiviral activity against influenza virus A (H1N1)

Dolastatin 10 (56)

Ulong Channel, Palau

Curacin A (55), D

Virgin islands

Cyanobacterium Lyngbya majuscula Cyanobacterium Symploca sp. VP642 Cyanobacterium Stigonema sp. Aspergillus sp. CNC-139

Anticancer activity through inhibition of the invasion of murine carcinoma colon 26-L5 cells Potent antifouling, antifungal activity

[137]

[136]

Cancer prophylaxis through induction of quinone reductase [135]

[130, 131]

[129]

[128]

[127]

[126]

Inhibition of the growth of human colon tumor cells with IC50 values of 0.68, 1.6, and 1.3 µg/mL for mixirin A, B, and C, respectively Antimitotic agent, inhibition of tubulin polymerization, potent inhibition of colcichine binding Selective toxicity against murine and human solid tumors

Mixirins A (54)–C (cyclic peptides)

Sea mud near the Arctic pole

Bacillus sp. MIX-62

Reference

Activity

Bioactive product

Location

Microbial isolate

Tab. 1.4 (continued)

16   1 Extremophiles: a promising source of novel natural products

1.5 Marine extremophiles 

 17

clinical trials due to their strong cytotoxicity [109]. The EU registered drug trabectedin (44) (Yondelis, anticancer) originates from a tunicate Ecteinascidia turbinate found in the Carribean and Mediterranean seas [3, 105] (Fig. 1.4). Marine environments have thus become favored targets for screening of bioactive products and constitute an intense area of research. For detailed information on the progress made in this area, see the various reviews [3, 110–115]. A database, MarinLit, established in 1970, is dedicated to marine natural products research (pubs.rsc.org/­marinlit/). Tab. 1.4 provides selected examples of the bioactive products (some structures (45–63) in Fig. 1.4) produced by the cold-water marine bacteria, Cyanobacteria and fungi. As in other environments, Actinobacteria constitute the most important group of eubacteria even in the marine environment, represented by several genera like Salinispora, Marinispora, Verriucosispora, etc. besides the omnipresent Streptomyces, producing promising bioactive compounds [41, 138, 139]. Genomes of several actinobacterial isolates from Arctic Ocean have shown the presence of at least two gene clusters involved in the synthesis of secondary metabolites [140]. Some novel compounds of interest isolated from marine actinomycetes include Abyssomycin C (46), a polycyclic polyketide antibiotic from Verrucosispora sp., diazepinomicin (ECO-4601) (47), farnesylated dibenzodiazepinone from Micromonospora sp., salinosporamide A (50), and a β-lactone-γ-lactam from the fermentation broth of Salinispora tropica (Tab. 1.4). Salinispora species were discovered to be an unusual group of Ascomycetes that required seawater for growth and 80% of these organisms produced culture extracts that inhibited in vitro growth of human colon carcinoma HCT-116. Among the initial reports on screening of deep sea microbiota for natural products, a family of novel macrolides, macrolactins A–F, were extracted from an unidentified Gram-positive bacterium isolated from the North Pacific [124]. The parent compound macrolactin A (53) exhibited selective antibacterial activity, inhibition of mammalian Herpes simplex viruses, and of melanoma cancer cells. Subsequently, different macrolactin isoforms and modified macrolactins have been discovered from marine Bacillus sp. cultures in several studies with spectrum of antibacterial, antifungal, and inhibitory activity on cytokine expression [141–143]. Cytotoxic cyclic peptides, mixirins (e.g. Mixirin A, 54) belonging to iturin-class acylpeptides, are among other bioactive products of interest found in marine Bacillus sp. MIX-62 isolated from a sea mud sample (Tab. 1.4). Marine Cyanobacteria have turned out to be an important source of metabolites, e.g. curacins (e.g. Curacin A, 55), dolastatin 10 (56), etc. with potential health benefits (Tab. 1.4). Dolastatin 10, the potent antitumor agent, was isolated originally in low yields from the invertebrate sea hare Dolabella auricu­ laria before being discovered in the Cyanobacteria Symploca sp. VP642 isolated from Hawaii [128]. As in other ecosystems, the marine fungal isolates are mainly represented by Penicillium sp., which have been found to be sources of cytotoxic metabolites, some examples of which are given in Tab. 1.4. A recent report further confirmed the predominance of Penicillium species among the culturable mycobiota isolated from marine segments at different depths [153–1463 m] in Antarctica Ocean, which

18 

 1 Extremophiles: a promising source of novel natural products

included Penicillium allii-sativi, Penicillium chrysogenum, Penicillium palitans, Pen­ icillium solitum besides Acremonium fusidioides, and Pseudogymnoascus verrucosus [144]. At least one isolate of each species exhibited antifungal, trypanocidal, leishmanicidal, antimalarial, nematocidal, or herbicidal activities. While a large number of bioactive products are being continuously reported from marine invertebrates including those mentioned above, majority of them are being shown or are believed to have their origins in the associated microalgae, Cyanobacteria, fungi, and heterotrophic bacteria [145, 146]. Bryostatins, a family of complex polyketides produced by a bryozoan, Bugula neritina, that are undergoing clinical trials as an anticancer drug based on their inhibitory effect on protein kinase C, have long been suspected to be produced by symbiotic bacteria [146]. The uncultivated gamma-proteobacterial symbiont “Candidatus Endobugula sertula” is apparently the source of bryostatin 1 (63) [147]. A series of diketopiperazines including a new natural product and two known phenazine alkaloid antibiotics were isolated from the culture broth of a sponge (Isodictya setifera)-associated bacterial strain of Pseudomonas aeruginosa collected at a depth of 30–40 m from Hut Point and Danger Slopes on Ross island, Antarctica [148]. A class I lantibiotic, subtilomycin with a broad antibacterial activity, has been reported from a Bacillus subtilis strain MMA7 isolated from the sponge Haliclona simulans collected on the west coast of Ireland [149]. Sponge-associated Actinomycetes isolates from Bay of Bengal, identified as Streptomyces sp., Saccharomonospora sp., and Micromonospora sp., have shown potent antibacterial activity against human pathogens [150]. In a study involving screening of 132 bacterial isolates from three Antarctic sponges for antimicrobial activity against 70 opportunistic pathogens, most were able to completely inhibit the growth of Burkholderia cepacia complex (Bcc) representing one of the most important cystic fibrosis pathogens [151]. The activity was attributed to an array of volatile organic compounds produced by the most active isolates, Pseudoalteromonas, Shewanella, and Psychrobacter. Fungi associated with Antarctic marine sponges, belonging to genera Geomy­ ces, Epicoccum, Penicillium, and Clodosporium possess antibacterial activity against several bacteria, while fungi associated with Antarctic macroalgae including Peni­ cillium, Palmaria, and Monostroma sp. displayed selective antifungal activity but no antibacterial activity [152, 153]. Penicillium steckii 6012 extract also caused inhibition of yellow fever virus [153]. Plinabulin (NPI-2358) (58), a synthetic analog of halimide obtained from Aspergillus sp. cultured from marine algae Halimeda lacrimosa, is being developed as an antitumor agent (Tab. 1.4).

1.6 Acidophiles Acidic environments, formed as a result of geochemical processes, e.g. acidic sulfur springs, and anthropogenic activities like coal and metal mining, invariably involve oxidation of elemental sulfur and sulfide minerals by molecular oxygen or ferric iron

1.6 Acidophiles 

 19

to produce sulfuric acid. They constitute an important source for acidophilic microorganisms; extreme acidophiles live in environments at pH 80,000 microbial natural products that have been reported to date, almost half (47%) have been shown to exhibit some form of biological activity [6], with natural products or their derivatives accounting for >40% of all Food and Drug Administration (FDA)–approved medications since 2000 [8, 9]. In certain categories of drugs, natural products dominate represent almost half of all drugs in the clinic. For example, 49% of anticancer drugs approved for clinical use between 1940 and 2014 were derived from natural products [8]. Since 2000, 77% of all FDA-approved antibiotics have been based on, or derived from, natural product scaffolds, of which all are of microbial origin [9]; this is in the context that almost 70% of currently prescribed antibiotics hark back to the “golden age” of discovery [10, 11]. Focusing on polyketide metabolites alone, of 7000 known structures, 20 have made it to the clinic [12]. This successes rate of 0.3% compares very favorably to 1000 atmospheres [18]. Marine microbes are highly abundant, with the world’s oceans predicted to contain on the order of 1029 individual microbial cells [19]. Estimates of the diversity of this marine life range from a few thousand species [20] to more than two million distinct taxa [21].

2.4 Life on the limit: how environmental conditions drive metabolic innovation The underlying biological mechanisms that organisms use to adapt to extreme environments provides a unique insight into the limits and fundamentals of cell biology. Understanding the expanding boundaries of macromolecular stability or the genetic code that constructs cellular molecules with the ability to function in one or more extreme conditions [22–28]. Although much still remains to be learned about the strategies employed for survival under such conditions, what is known to us is that these microorganisms utilize novel biochemical pathways. They provide much needed alternatives to liable mesophilic molecules that lack stability and activity in more extreme conditions. This is most evident for extremophilic enzymes. There is significant biotechnological potential for enzymes with the ability to remain catalytically active at extreme temperatures, at acidic or basic conditions, at high salinity or those able to retain activity in organic solvents. These properties make them attractive targets for use in industrial applications [29]. The biosynthetic potential of extremophiles, which are often difficult to isolate and study, is highly unexplored. Targeting these microorganisms as a novel source of natural product-based drug leads therefore represents an attractive route to mitigating the risk of rediscovering known natural products from microbial metabolomes. Researchers have therefore, in recent years, turned their attention to the isolation and characterization of extremophile microorganisms, and a number of success stories are now beginning to emerge. Here, we outline some notable examples of pharmaceutical leads identified from extremophile natural products that show promise for use in the treatment of a range of human and animal diseases.

2.5 Lomaiviticins During a search for natural products with anticancer properties, McDonald et al. isolated the natural product compound namenamicin from the marine ascidian

2.5 Lomaiviticins 

 47

(sea squirt) Polysyncraton lithostrotum, which was collected off the coast of Fiji [30]. Fig. 2.2 shows the chemical structure of namenamicin (6), which was shown to be a member of the enediyne family of antitumor natural products (6–8). These had previously been isolated exclusively from actinomycete bacteria [31] and so it was suggested that namenamicin was also of microbial origin. Attempts were made to isolate this supposed microorganism from the invertebrate [32]. A phage lambda-based biochemical assay [33] was used to detect, identify, and purify DNA-damaging agents in the fermentation broth of cultured bacteria that were associated with the ascidian, and the halophile strain LL-371366 emerged as a promising candidate for enediyne production. This halophilic bacterium was originally classified as a new species, Micromonospora lomaivitiensis, but later phylogenetic analysis [34] revealed it to be a strain of Salinispora pacifica, an obligate marine actinomycete [35]. In addition to namenamicin, two novel glycosylated diazo benzofluorene dimers were also isolated from strain LL-371366 [32] and named lomaiviticin A and B (9, 10) (Fig. 2.2). Lomaiviticin A was tested for cytotoxicity against a panel of 24 cancer cell lines and found to be extremely potent, with a half maximal inhibitory concentration (IC50) down to 7 pM in some of the cell lines [32, 36]. The cytotoxicity profile of lomaiviticin A in these assays was unlike any other known DNA-damaging anticancer drug [32], implying that its mode of action was unique. Lomaiviticins A and B were also both found to have antibacterial activity in plate assays [32]. The diazotetrahydrobenzo[b]fluorene moiety that forms the core of the lomaiviticins is highly unusual, reinforcing the idea that extremophiles will harbor unfamiliar metabolites. This tetracycle consists of a napthoquinone fused to a diazocyclopentadiene, which is in turn bonded to an oxidized cyclohexene ring, resulting in distinctive extended pi-conjugation. Diazo functions are rare in natural products, and little is known about their biosynthesis [37]. The only other metabolites known to share the diazofluorene moiety are the kinamycins (11a–f), which were isolated from soil bacteria in Japan (Fig. 2.2) [38]. However, key structural differences differentiate these molecules. Kinamycins possess a single diazofluorene, whereas lomaiviticins contain two such residues linked via a carbon-carbon bond into a C2-symmetric dimer. Kinamycins are less oxidized than lomaiviticins and are not functionalized by sugar residues. These differences appear critically important for activity, as kinamycins are two orders of magnitude less cytotoxic than lomaiviticin A [39]. Attempts to achieve the total chemical synthesis of lomaiviticins have been impeded by two obstacles [40]. First, forming the carbon-carbon bond that links the two diazofluorene monomers with controlled stereochemistry is difficult due to the steric congestion around the dimer interface, which can impose unpredictable destabilizing interactions [41]. Secondly, the β-hydroxy ketone at the dimer interface readily undergoes elimination and aromatization reactions, resulting in a loss of stereospecificity. Although impressive progress has been made in synthesizing substructures of lomaiviticins [41, 42], including the core carbon skeleton of the molecule but lacking the sugar residues [43], complete synthesis of these natural products has not yet been reported. It has also proven challenging to isolate significant ­quantities of

48 

 2 The extremophilic pharmacy: drug discovery at the limits of life

Fig. 2.2: Chemical structures of namenamicin (6) and closely related enediyne natural products (7, 8), lomaiviticins (9, 10), kinamycins (11a–f), and bleomycin (12).

lomaiviticin A from fermentation broths. However, Woo et al. established that another bacterium, Salinispora tropica produces high levels of lomaiviticin C during fermentation [44]. This metabolite is identical to lomaiviticin A, with the exception of the elimination of the diazo group on one of its diazofluorene moieties. The r­ esearchers

2.5 Lomaiviticins 

 49

developed a simple semisynthesis of lomaiviticin A from this more abundant natural product, allowing detailed studies on the activity of lomaiviticin A in parallel with lomaiviticin C and kinamycin C [39]. Kersten and coworkers identified the lomaiviticin biosynthetic cluster in Salinispora tropica [45]. Designated the lom cluster, this region was predicted to encode 59 open reading frames. All of the genes thought to be involved in the biosynthesis of the diazofluorene core were identified, along with two glycosyltransferases consistent with the observed glycosylation pattern of the lomaiviticins. Significantly, a putative flavin adenine dinucleotide-dependent monooxygenase gene (Lom19) shows 76% sequence identity to ActVA-Orf4 from the Streptomyces coelicolor A3(2) actinorhodin pathway [46]. This enzyme catalyzes carbon-carbon bond formation resulting in the dimerization of a multicyclic aromatic molecule, which suggests that Lom19 is the diazofluorene dimerase. The lomaiviticin cluster was later identified in Salinispora pacifica, and candidate enzymes involved in diazo synthesis and diazofluorene dimerization were proposed by comparison with kinamycin clusters [34]. Characterization of the enzymes involved in lomaiviticin biosynthesis, in particular the dimerase and diazo synthases, is expected to provide new insight and novel tools for biocatalysis. Metabolic engineering of the lom cluster to produce new lomaiviticin analogs with improved properties is also an important goal. The major activity of the lomaiviticin family appears to be introducing nicks into DNA. Lomaiviticin A nicks double-stranded DNA (dsDNA) and induces double-strand breaks (DSBs) in the presence of a reducing agent [39]. The ratio of single-strand breaks (SSBs) to DSBs induced by lomaiviticin A (5:1) was much lower than expected if DSBs arose due to the cumulative effects of unrelated SSBs; this ratio is comparable to that of bleomycin (12) (6:1), which is known to effect two SSB events on complementary strands of dsDNA without dissociating [47]. Taken together, this evidence supports a model in which lomaiviticin A binds dsDNA and nicks each strand of the duplex in a stepwise manner, producing extremely cytotoxic DSBs [48, 49]. The inability of lomaiviticin C and kinamycin C to do the same implies that both diazofluorene groups are required for this activity. Subsequent studies by the same group established that vinyl radicals of lomaiviticin A can form by nucleophilic addition to the diazo moieties [50]. These radicals are capable of abstracting hydrogen atoms from methanol and acetone. These data were used to propose a model in which an unidentified nucleophile (possibly DNA itself) forms an adduct at the first diazo group, triggering the formation of the first vinyl radical, which removes a hydrogen atom from the DNA backbone of one strand. This leads to a SSB by a well-understood mechanism [51], and the process is repeated with the second diazo moiety and complementary DNA strand to produce a DSB. Unexpectedly, the second nucleophilic addition was found to be far slower than the first, offering an explanation for the 5:1 ratio of SSBs to DSBs [50]. This mechanistic model implies that lomaiviticin A adopts a DNA-bound conformation in which both diazofluorenes are simultaneously close to each dsDNA strand. This was confirmed by a combination of proton nuclear magnetic resonance spectroscopy of

50 

 2 The extremophilic pharmacy: drug discovery at the limits of life

a palindromic DNA duplex bound to lomaiviticin A and molecular modeling of the complex based on NMR-derived constraints [52]. The structure reveals that both diazofluorene groups are inserted into the minor groove of the DNA, forcing three bases out of the duplex. The energetic cost of this base flipping may be ameliorated by the favorable base-stacking interactions of the diazofluorenes and remaining DNA bases and by electrostatic interactions between the amino sugar and the phosphate backbone. This binding mode places the diazo carbons of lomaiviticin A close enough to key deoxyribosyl hydrogen atoms for abstraction and strand scission. The model also accounts for the 10–100-fold reduced potency of lomaiviticin B, as its two additional ether linkages between diazofluorenes are expected to prevent it adopting the same mode of binding DNA. The diazofluorenes are a fascinating target for further research, with the potential to provide new insights in synthetic chemistry, enzymology, and cancer biology. Lomaiviticin A is being investigated as a monotherapy and combination therapy for DSB repair-deficient tumors [49, 53].

2.6 Salinosporamide A The halophile actinomycete Salinispora tropica is also the source of one of the most significant discoveries in extremophile natural product research in recent years: salinosporamide A (13) [54–58]. Screening of organic extracts of cultured Salinispora strains by Feling et al. demonstrated a high rate of antibiotic and anticancer activity [59]. The crude acetone fraction of Salinispora strain CNB-392 exhibited highly cytoto­ xic activity against a human carcinoma cell line in vitro, with IC50 values of 80 ng/mL-1. The active ingredient was identified as salinosporamide A (Fig. 2.3), which contains a γ-lactam-β-lactone bicyclic core shared with Omuralide (14), a compound produced by a terrestrial Streptomyces species [59, 60]. Salinosporamide A is elaborated by the addition of a methyl group at the C3 ring juncture, a chloroethyl group at C2, and a cyclohexene at the C5 position. Omuralide was a milestone in cancer treatment as the first truly specific inhibitor of the proteasome, a catalytic complex responsible for nonlysosomal proteolysis in the cell [61–65]. The modifications found in salinosporamide A caused the

Fig. 2.3: Chemical structures of salinosporamides A and B (13, 15), and omuralide (14).

2.6 Salinosporamide A 

 51

inhibition of proteasome activity with a potency 35 times greater than omuralide [59]. S ­ tructural biology revealed how salinosporamide A bound to the 20S proteasome subunit of yeast. The six catalytic subunits present in the proteasome contain an N-terminal nucleophilic threonine residue, which is essential for polypeptide hydrolysis and forms the target for a covalent bond with omuralide, affording the mechanism of inhibition [66]. It was thus postulated that the ester linkage between the Thr10 γ-hydroxyl and the β-lactone carbonyl would form an equivalent adduct in salinosporamide A. However, although nucleophilic addition of the β-lactone ring does occur, upon ring opening, the C3 hydroxyl displaces a key water molecule in the enzyme active site, hindering deacylation and forming a cyclic ether with the chloroethyl group of salinosporamide A. The resulting protonated threonine deactivates the catalytic N-terminus [67]. The presence of the chlorine atom is thus vital for the potency of salinosporamide A and indeed salinosporamide B (15), a deschloro analogue of salinosporamide A, is 500 times less potent than its chlorinated counterpart [68, 69]. The general relevance of halogen atoms in pharmaceuticals has long been recognized, and marine environments represent an excellent source of novel halogenation pathways [68]. It is clear that the ability of marine extremophiles to produce halogenated natural products represents a significant resource for promising drug leads. The biosynthetic route to both salinosporamide A and B was first characterized by Moore and coworkers [69–71], who proposed the involvement of ­chloroethylmalonyl-coenzyme A (coA) (16) in the addition of the unique polyketide extender unit chlorobutyrate (Fig. 2.4). This halogenated precursor was shown to be formed by an S-adenosyl-L-methonine (SAM, 17) dependent chlorinase, a new family of halogenase enzymes that utilize a rarely observed nucleophilic substitution strategy, converting SAM to 5’-chloro-5’deoxyadenosine (5’-CIDA, 18) [69]. Production of salinosporamide A from the producer microorganism was hampered by the unsuitability of large-scale fermentation vessels for saline fermentation due to corrosive effects and the inherent aqueous instability of the β-lactone ring. This was overcome by the addition of a solid resin able to bind to the active product, increasing production 69-fold [72]. Salinosporamide A was the first example of the use of saline fermentation to produce clinical trial material, paving the way for future natural products to be produced via the fermentation of halophilic bacteria [73]. Total chemical synthesis was achieved only a year after salinosporamide A’s chemical structure was first published [74, 75]. This synthesis is highly stereo-controlled and enantioselective, requiring the formation of five stereocenters within a fused γ-lactam-β-lactone bicyclic core. Salinosporamide A was licensed by Nereus Pharmaceuticals and is now showing great promise in clinical trials under the trade name Marizomib [76, 77]. The rise of salinosporamide A from the sea sediment to the clinic is an excellent example of the potential of the extremophile pharmacy (Fig. 2.5).

Fig. 2.4: Proposed biosynthetic pathway to chloroethylmalonyl-CoA (16) as a polyketide synthase extender unit in salinosporamide A (13) biosynthesis.

52   2 The extremophilic pharmacy: drug discovery at the limits of life

2.7 Marinostatin 

 53

2005: IND filed Preclinical development Preclinical models API Manufacturing Formulation Toxicology

2006: Phase 1 Solid tumors and lymphomas

2007: Phase 1 Multiple myeloma

Discovery and development of Salinosporamide A 2002: Salinispora discovered

2006: Crystal structure 2007: Total in complex with 20S synthesis proteasome 2007: Salinispora 2003: Salinosporamide A isolated, 2005: Efficacy in mouse tropica genome structure elucidated, mode of 2006: Efficacy in mouse multiple myeloma sequenced action and cytotoxicity established colon cancer xenograft xenograft models models 2004 and 2005: Total synthesis complete

Fig. 2.5: Discovery and clinical development of salinosporamide A. Adapted from Fenical et al. (2009).

2.7 Marinostatin Proteases are an abundant enzyme class involved in diverse and critically important biological processes. Proteolysis is required for cell cycle transitions, apoptosis, immune regulation, and cell signalling; accordingly, proteases are implicated in the pathogeneses of many diseases, including inflammatory disorders, cancer, and infections [78]. Natural products that can act as protease inhibitors are highly valued both as potential therapeutics and as tools for basic science. Imada et al. [79] used an agar diffusion method to screen for protease inhibitors in bacteria from seawater and sediment samples from sites around Japan. Of the nearly 2000 isolates screened, three inhibited the hydrolysis of casein by the protease subtilisin. A strain termed B-10-31 produced the largest inhibitory zone and was selected for further study. This strain was eventually identified as Pseudoalteromonas sagamiensis, a halophile marine ­bacteria [80]. The protease inhibitor produced by P. sagamiensis was determined to be a 12-residue peptide [81]. This peptide, termed marinostatin (26), was the first peptide protease inhibitor isolated from a marine microorganism and is the smallest natural product member of the serine protease inhibitor family [82] (Fig. 2.6). Early evidence indicated that marinostatin specifically inhibited serine proteases (with the exception of trypsin) and that this inhibition could be the result of unusual ester linkages [81, 83]. Subsequent analyses provided a detailed model of marinostatin inhibition, which we describe below. The interactions of proteases and their protein inhibitors have been extensively reviewed [84, 85]. Canonical serine protease inhibitors bind the enzyme in a

54 

 2 The extremophilic pharmacy: drug discovery at the limits of life

Fig. 2.6: Chemical structure and schematic representation of marinostatin (26).

substrate-like manner, with inhibition thought to occur due to the inhibitor being “too perfect” a substrate. The enzyme-inhibitor complex forms a classic lock-and-key interaction, with an optimal fit mutually stabilized by hydrogen bonds, electrostatic interactions, and van der Waals forces. The inhibition constant (Ki) values of serine protease inhibitors are many orders of magnitude lower than the dissociation constant (KD) values of their natural substrates. The residues of protease substrates and inhibitors are numbered according to distance from the scissile peptide bond between P1 and P1′, as shown in Fig. 2.7. The identity of residue P1 is critical for binding and determines the specificity of the interaction. Different serine proteases have different amino acid preferences at this position, with proline often required within two positions of the reactive bond to enforce correct geometry. Most serine protease inhibitors have a flexible binding loop containing the scissile bond, and a compact hydrophobic core stabilized by disulphide bridges, which were thought to be absolutely required for inhibition [85]. However, the twelve residues of marinostatin are too few in number to pack into a hydrophobic core, and its sequence lacks cysteines. Kanaori et al. determined the solution structure of marinostatin by 1H NMR spectroscopy and found that it could be superimposed on the structure of OMTKY3 (turkey ovomucoid third domain), a canonical serine protease inhibitor bound to subtilisin [86], and shared similar Ramachandran angles over the binding loop [87]. Further work revealed how marinostatin achieves the structural rigidity required of canonical serine protease inhibitors. The peptide contains two ester linkages between the beta-hydroxyl and beta-carbonyl groups of Thr3-Asp9 and Ser8-Asp11, which give it a bicyclic structure. The carbonyl of the 3-9 linkage is involved in a hydrogen bond with the backbone amide of Arg5, as evidenced by the greatly reduced hydrogen-deuterium exchange rate of this proton. This hydrogen bond is thought to suppress the conformational fluctuation of marinostatin, helping to protect the reactive peptide bond from cleavage, and is essential for inhibitory activity. An analogue of marinostatin with cysteine residues at positions 3 and 9 forms a stabilizing disulphide bridge but is significantly more flexible due to the loss of this

2.7 Marinostatin 

 55

Fig. 2.7: General mechanism of polypeptide cleavage by a protease. The scissile peptide bond is shown in red. Substrate amino acid residues are labeled as P1, P1′, P2, P2′, and P3, P3′. Corresponding enzyme binding sites are labeled as S1, S1′, S2, S2′, S3, S3′.

key hydrogen bond. Consequently, it can only inhibit protease activity temporarily before its proteolytic inactivation [88]. Further structure-activity studies shed light on the role of Pro7, which adopts a cis conformation to force the marinostatin backbone into a rigid beta-turn that promotes formation of the ester-amide hydrogen bond [89]. Forcing this region into the trans conformation by replacing Pro7 with a trans-olefin abolished inhibitory activity and distorted the structure of marinostatin [82]. The N-terminal Phe1 residue of marinostatin was also shown to be essential for activity due to predicted binding pocket interactions [89]. Thus, it can be seen that mutually stabilizing interactions between serine proteases and marinostatin confer rigidity to the inhibitor, ensure strong binding, and prevent deformation of the scissile peptide bond, protecting it from hydrolysis [85]. Recently a total synthesis of marinostatin was achieved by sequential esterification of the linear peptide using orthogonal protecting groups [90]. This work paved the way for the rational design of peptide analogues with different protease targets. The simplicity of marinostatin makes it an attractive target for modification. It is the smallest canonical protease inhibitor in nature [87] and the first nine residues comprise the minimal sequence required for activity [89]. The specificity of marinostatin was successfully changed from subtilisin to trypsin by swapping residues four and five, and this modified peptide inhibited trypsin more potently than leupeptin, which is commonly used to prevent proteolytic degradation by endogenous proteases during recombinant protein purification [89]. The marinostatin biosynthetic gene cluster was first identified in 1998 [91], and since then, it has become clear that marinostatin is a member of the ribosomally synthesized and post-translationally modified peptide (RiPP) family of natural products [92]. RiPP genes are ubiquitous in microbial genomes. The immediate translation product is a precursor peptide that contains a leader sequence for recognition by biosynthetic enzymes. This precursor is post-translationally modified and then undergoes proteolysis to remove the leader sequence before export from the cell. This pathway makes RiPPs prime candidates for metabolic engineering [93]. The biosynthetic enzymes recognize the leader but are permissive of varying downstream sequences and the biosynthesis relies on only a small number of enzymes, ­simplifying

56 

 2 The extremophilic pharmacy: drug discovery at the limits of life

heterologous expression. Marinostatin belongs to a subgroup of RiPPs known as the microviridin family. The biosynthesis of microviridins is now well understood [94–96] and has been reconstituted in vitro, allowing for the generation of diverse and highly relevant protease-targeted peptide inhibitor libraries [97]. Activity-based probes have recently emerged as enabling technologies for dissecting protease functions and identifying new drug targets [98–101]. These small molecules consist of a recognition element that specifically binds a target protease, an electrophilic “warhead” that reacts irreversibly with the protease active site to form a covalent adduct, and a tag that can be used to purify or track the probe by fluorescence or binding to a partner. Activity-based probes have been extensively applied to the study of biological systems. In combination with super-resolution microscopy, these probes can noninvasively track single protease molecules in real time, allowing the dynamics and spatial organization of protease activity to be visualized [102, 103]. Activity-based probes are particularly useful for systems in which genetic tools are not well developed. Selective probes have been designed for Mycobacterium tuberculosis proteases to study their role in virulence and monitor infections [104], and activity-based probes have been used to identify macrophages as the cells that express cathepsin proteases in response to interleukin 4 secreted by cancer cells, promoting tumor growth and invasion [105]. These specific tool compounds increase our understanding of complex biological processes and identify potential drug targets. As a simple and specific protease inhibitor with a well-characterized structure, mechanism, and biosynthetic pathway, marinostatin is not only an exciting target for therapeutic development but also a valuable starting point for the development of future biochemical probes.

2.8 Abyssomicin C The marine environment has also served as a source of novel antibiotics. This is demonstrated by the discovery of the abyssomicins, which act as inhibitors of tetrahydrofolate (THF) biosynthesis. THF is required for several key metabolic reactions in bacteria, including the biosynthesis of amino acids and purine bases [106, 107] (Fig. 2.8). The production of THF is thus an essential pathway in many microorganisms but is absent in higher organisms, which instead obtain THF from their diet [107]. This makes THF biosynthesis an attractive and well-established antibiotic target [106, 107]. Bister et al. focused on a particular step in THF biosynthesis: the production of the precursor para-aminobenzoate (pABA; 27) (Fig. 2.8). They isolated 201 extremophile actinomycetes from deep sea sediment from the Sea of Japan and screened extracts from these bacteria for the ability to specifically inhibit the biosynthesis of pABA [108]. This approach yielded a new compound abyssomicin C (32) from the halophile Verrucosispora strain AB 18-032 (Fig. 2.9). Abyssomicin C was found to inhibit the

2.8 Abyssomicin C 

 57

growth of pathogenic strains of Staphylococcus aureus and Mycobacterium tuberculosis with minimal inhibitory concentration values in the low micromolar range but did not affect Gram-negative bacteria [108–110]. The complex structure of abyssomicin C can be broken down into three structural motifs, as shown in Fig. 2.9. These are (i) an oxabicyclo[2.2.2]octane core spirolinked to (ii) a tetronic acid ring, fused to (iii) an 11-membered ring including an α, β-unsaturated ketone. The similarity of the oxabicyclooctane system to a solution conformation of chorismate suggested that abyssomicin C might act as a substrate mimic for the enzyme 4-amino-4-deoxychorismate (ADC) synthase [108] (Fig. 2.8). Additional members of the abyssomicin family (33–36) (Fig. 2.9) were later discovered in related species of bacteria. Structure-activity relationship studies aided by various total syntheses of the abyssomicins helped to define the mechanism of antibiotic activity [111–115]. An enone group at position C7-C9 was found to be essential for antibiotic activity. This enone acts as a Michael acceptor, which inhibits ADC synthase by covalently and irreversibly binding to a catalytic cysteine residue [108, 109]. Further structure-activity relationship studies with synthetic analogues of abys­ somicin C have established that the C11 hydroxyl and C3 carbonyl groups are not required for activity; that C11 benzyl ethers are more potent than the natural product; and that removing the three methyl groups lowers cytotoxicity by three orders of magnitude [116, 117]. However, the nonspecific reactivity of the enone group remains problematic, and desmethyl abyssomicin C remains cytotoxic at antibacterial concentrations [117]. An interesting exception within this family is abyssomicin J (38), which lacks the Michael acceptor but retains antibiotic activity. It appears that this thioether-linked dimer undergoes a reverse Michael reaction in situ to generate active

Fig. 2.8: Biosynthetic route to the THF precursor pABA (27).

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Fig. 2.9: Chemical structures of selected abyssomicins (32–36) and cisoid and transoid enones.

abyssomicin C, which would explain why its MIC value is half that of abyssomicin C [114]. Delivering abyssomicin C in this form as a prodrug could enhance its bioavailability by stabilizing the reactive enone. During their total synthesis of abyssomicin C, Nicolaou and colleagues discovered that this molecule exhibits atropisomerism [111]. Atropisomers are stereoisomers that arise due to restricted rotation around a single bond, with an energy barrier of at least 23.3 kcal/mol [118]. The highly strained 11-membered ring of abyssomicin C restricts the rotation of the C7-C8 bond, so that the C7 carbonyl is either in a transoid (abyssomicin C) or cisoid (atrop-abyssomicin C) conformation. Nicolaou et al. found that atrop-abyssomicin C is 1.5 times more potent an antibiotic than its atropisomer and compared the X-ray crystal structures of the two compounds to propose a

2.8 Abyssomicin C 

 59

mechanism for this difference [115]. The O=C7-C8=C9 dihedral angle of abyssomicin C is 144.8°, whereas in atrop-abyssomicin C, it is 26.4° (Fig. 2.9). These two electron rich systems are forced closer together in the latter, increasing the degree of conjugation and making the enone a better Michael acceptor, which was later confirmed in kinetic experiments [111]. The atropisomers can interconvert in the presence of acid, which explains why atrop-abyssomicin C was not identified in early studies that used acidic solvents for high performance liquid chromatography (HPLC). A later study that avoided the use of acidic HPLC solvents found that atrop-abyssomicin C is the major biosynthetic product of fermentation [113]. The biosynthetic gene cluster for the abyssomicins (aby) was identified in Verrucosispora maris [119–121]. Abyssomicins belong to the spirotetronate polyketide family of microbial metabolites, defined by a cyclohexene ring spiro-linked to a tetronic acid group [122]. The expected PKS and tetronate biosynthesis genes were identified, along with various putative oxygenases, regulators, and exporters. Feeding experiments with radiolabeled precursors demonstrated that the linear abyssomicin C polyketide is built from acetates, propionates, and an unidentified glycolytic metabolite. The authors proposed a biosynthetic scheme including a [4 + 2] cycloaddition to form the spiro-link but could not identify a candidate enzyme for the transformation. Later studies on the related spirotetronate versipelostatin identified a small enzyme VstJ, which catalyzed a stereoselective [4 + 2] cycloaddition and found a small homologous 429 bp open reading frame in the aby cluster, which has not previously been recognized as a gene [123]. The predicted protein product of this gene, later named AbyU, was identical to a 141-residue protein from another V. maris strain in which abyssomicin production had not been reported [124]. The Diels-Alder reaction is a [4 + 2] cycloaddition reaction that involves concerted reorganization of a six-electron system to form a cyclohexene [125]. Most enzymecatalyzed reactions involve either ionic two-electron or radical one-electron processes, and pericyclic reactions are extremely rare. For a long time, it was unknown if any enzyme catalysts of Diels-Alder reactions (Diels-Alderases) existed [126]. While a small number of enzymes have been shown to catalyze [4 + 2] cycloadditions, whether they proceeded via a concerted Diels-Alder route with a single cyclic transition state was unknown. Biocatalysis of the Diels-Alder reaction is a major goal as it would allow new, more efficient, and more environmentally friendly routes to valuable bioactive compounds. Byrne et al. [127] confirmed that AbyU catalyzes [4 + 2] cycloadditions with substrate analogues and determined the crystal structure of the recombinant enzyme. These findings enabled the establishment of a formalized biosynthetic route to the compound (Fig. 2.10). AbyU was shown to be a homodimer comprising a pair of eight-stranded antiparallel beta barrels, each of which possesses a central hydrophobic channel forming the active site. Molecular dynamics simulations of the enzyme with substrate showed that AbyU catalyzed a bona fide Diels-Alder reaction. The active site has excellent complementarity to its natural substrate and holds the molecule in a reactive conformation from which the Diels-Alder reaction can occur with a lowered free energy barrier. The simplicity of AbyU makes it an attractive

Fig. 2.10: Biosynthesis of the abyssomicin C spirotetronate core. The linear polyketide 37 is condensed with a glyceryl unit to form 38. Subsequent enzyme catalyzed tailoring reactions give 40, the substrate for the AbyU catalyzed Diels-Alder reaction to give 41.

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2.9 Macrolactins 

 61

candidate for reengineering, via the mutation of key amino acid residues within the barrel core. It is hoped that such an approach could lead to the development of functionally optimized non-natural abyssomicins with improved clinical utility.

2.9 Macrolactins In 1989, as part of a research program to culture and characterize marine microorganisms, Gustafson et al. isolated a Gram-positive bacterium from a sediment core 980 m below sea level off the coast of San Francisco [128]. Using the biochemical methods available at the time, the group found the strain taxonomically undefinable, but noted that it was a halophile with a strong salt requirement for growth and likely a piezophile due to the hydrostatic pressure of the deep-sea environment. When grown in liquid culture, this bacterium produced a novel family of macrolides with 24-membered macrocyclic lactone rings named macrolactins (42–49) (Fig. 2.11). The identity and abundance of the macrolactins produced by this strain were dependent upon the culture conditions used, and six individual macrolactins were isolated [128]. Macrolactin A is considered the archetype of the family, with characteristic polyenes, a lactone linkage, three hydroxyl groups, and a single methyl substituent. The hydroxyl moieties are modified by dicarboxylic acids and sugars, and geometric isomerism occurs around the alkene groups to generate the structural diversity of the family. Later analyses established the absolute and relative stereochemistry of macrolactins, and a total chemical synthesis was established to compensate for the unreliable fermentation of the original producing strain [129, 130]. In the original study, macrolactin A was reported to possess an extraordinary range of bioactivities. It inhibited the growth of B. subtilis and S. aureus, murine melanoma cells, herpes simplex virus, and HIV replication at the microgram scale in in vitro assays [128]. Subsequent work established that macrolactin A could protect neurons from L-glutamate toxicity in a model of brain ischemia [131] and act as a biological pest control agent in potato scab disease [132]. Furthermore, macrolactins A and F were rediscovered in a screen for squalene synthase inhibitors and proposed as potential therapeutics for hyperlipidemia [133]. Other macrolactins were subsequently discovered in marine and soil Bacillus species, and to date, the family has expanded to include macrolactins A-W and their derivatives (42–49) (Fig. 2.11) [134–142]. Unlike the rest of the family, macrolactins S, V, and W exhibit antibacterial activity against both Gram-negative and Gram-positive bacteria. Macrolactin V exhibits a potent MIC of 0.1 µg ml-1 against E. coli, B. subtilis, and S. aureus. Its epimer macrolactin S has similar activity but with a different profile, being ineffective against B. subtilis [141]. Macrolactin W was much less active but could additionally inhibit the growth of P. aeruginosa, an opportunistic Gram-negative pathogen, which is a frequent cause of hospital-acquired infections [142, 143].

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Fig. 2.11: Chemical structures of selected macrolactins.

In the search for new antibiotics, the increasing number of sequenced bacterial genomes has allowed for rational selection of conserved bacterial genes as antibacterial targets. If a gene is essential in bacteria but no human homologue exists, it should in principle represent a good target for inhibition. Peptide deformylase (PDF), an essential bacterial enzyme that removes formate from the N-terminus of proteins, is one such target [144]. PDF inhibitors have reached clinical development [145, 146] but were discontinued due to issues with their selectivity and in vivo stability, which is a general issue with peptide therapeutics [147]. During a screen for novel PDF inhibitors, macrolactin N was isolated from a soil-dwelling B. subtilis strain and found to inhibit S. aureus PDF with an IC50 of 7.5 µM [137]. The same group identified

2.9 Macrolactins 

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macrolactins O-R as weaker PDF inhibitors and found that the PDF-inhibiting macrolactins have antibacterial activity against both Gram-negative and Gram-positive species [138]. Computational studies [147] showed that macrolactins probably bind to the same site as unrelated PDF inhibitors and that hydrophobic and van der Waals forces provide most of the binding energy. This provides a robust starting point for the rational redesign of macrolactins as antibiotics. Macrolactin A and its 7-O-succinyl and 7-O-malonyl derivatives (SMA and MMA, respectively) have attracted the most interest as potential therapeutics. In 2013, Young-Hoon et al. were granted a patent for the use of these three compounds as anti-inflammatory drugs, taking advantage of their ability to inhibit the production of proinflammatory cytokines and nitric oxide [148]. In 2014, they were also granted a patent for the use of macrolactin A derivatives as anti-angiogenic compounds [149]. It transpired that SMA could inhibit a cell signalling component common to inflammation and angiogenesis, phosphatidylinositide 3-kinase (PI3K) [150, 151]. SMA inhibits PI3K in vitro and suppresses phosphorylation of protein kinase B, a downstream PI3K target in vivo [152]. SMA strongly suppresses angiogenesis in a standard chick embryo assay and displays no cytotoxicity to human cells at concentrations up to 50 µM [151]. SMA has also demonstrated antitumor activity in mouse models of cancer and appears to act by inhibiting PI3K and tankyrase, a component of the Wnt pathway, which inappropriately promotes cell survival and proliferation in many cancers [153, 154]. SMA shows synergistic effects with common anticancer agents like cisplatin, allowing them to be used at lower doses to minimize side effects [155, 156]. An appropriate commencement dose for humans (~1.5 mg/kg) has been calculated for SMA [157], and the route toward clinical trials seems clear. The macrolactin biosynthetic cluster (mln) was identified in the soil bacterium Bacillus amyloliquefaciens and characterized as a trans-AT type I modular polyketide synthase (PKS) system [158]. This cluster has several unusual characteristics including split PKS modules, the absence of enoyl reductase and dehydratase domains predicted from the chemical structure of the macrolactins, and duplicated acyl-carrier protein domains, several of which lack the universally conserved serine residue essential for function. Feeding experiments with radiolabeled substrates indicated that acetate is the sole precursor of the macrolactin carbon skeleton, and in combination with the operon structure, these were used to propose a biosynthetic route to the compound. A key enzyme required for macrolactin biosynthesis is the glycosyltransferase designated BmmGT1 [159]. Intriguingly, this enzyme is not specific to macrolactin biosynthesis but also acts on the polyketide antibiotic bacillaene. Overexpression of BmmGT1 resulted in the production of two novel macrolactin A derivatives glycosylated at positions 13 and 15, and incubation of the recombinant enzyme with UDP-N-acetylglucosamine produced a further new derivative. The substrate promiscuity of BmmGT1 could be exploited to generate new macrolactin analogues with improved bioavailability or reduced toxicity. Zotchev et al. developed an approach

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for generating in silico libraries of macrolactins with predicted bioactivities to guide metabolic engineering efforts [160]. Their program, Bio-generator, indicates the PKS modifications required to produce the selected macrolactins and aims to reduce the laborious process of combinatorial biosynthesis of polyketides. The wealth of information about macrolactin biosynthesis should encourage rational redesign studies to harness the full potential of these remarkable metabolites [161].

2.10 Conclusions and future prospects The examples discussed above show how the novelty and diversity of extremophile natural products can provoke new directions in pharmaceutical development. Much of Earth’s microbial life remains uncharacterized and there is clear potential for new discoveries from these microorganisms [162–165]. There is plenty of historic precedent to suggest that the complexity and biological activity of these natural products will be beyond both our imagination and synthetic capability. The major barrier to progress, both historically and in the present day, remains the need to screen cultivated bacteria for the production of functional natural products. This has proven limiting since the majority of microbes apparently resist laboratory culture. However, new technologies that are now emerging may help to overcome this challenge or even obviate the need for cultivation altogether. Recent advances in DNA sequencing and synthesis methods are now driving genomics-based natural product drug discovery from cultured and uncultured microorganisms. These approaches are becoming ever more accessible and affordable and are underpinned by a well-developed bioinformatics toolkit. Advances in synthetic biology are facilitating the reconstitution of natural product pathways in model or industrial hosts, enabling the production and rapid characterization of natural products from extremophiles at scale. Such approaches can increasingly be undertaken using liquidhandling robotics, allowing natural product libraries to be screened in a high-throughput fashion. This is being supported by the increasing sensitivity, sophistication, and automation of analytical methods and separation technologies. Extremophiles undoubtedly represent the planet’s largest unexplored pool of biological and chemical novelty [166]. Through continued characterization and interrogation of this resource, it is hoped that it will be possible to identify the pharmaceuticals of the future.

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[72] Tsueng G, Lam KS. Stabilization effect of resin on the production of potent proteasome inhibitor NPI-0052 during submerged fermentation of Salinispora tropica. J Antibiot. 2007;60(7):469–72. [73] Fenical W, Jensen PR, Palladino MA, Lam KS, Lloyd GK, Potts BC. Discovery and development of the anticancer agent salinosporamide A (NPI-0052). Bioorg Med Chem. 2009;17(6):2175–80. [74] Endo A, Danishefsky SJ. Total synthesis of salinosporamide A. J Am Chem Soc. 2005;127(23):8298–9. [75] Reddy LR, Saravanan P, Corey EJ. A simple stereo-controlled synthesis of salinosporamide A. J Am Chem Soc. 2004;126(20):6230–1. [76] Potts BC, Albitar MX, Anderson KC, et al. Marizomib, a proteasome inhibitor for all seasons: preclinical profile and a framework for clinical trials. Curr Cancer Drug Targets. 2011;11(3):254–84. [77] Di K, Lloyd GK, Abraham V, et al. Marizomib activity as a single agent in malignant gliomas: ability to cross the blood-brain barrier. Neuro-Oncology. 2016;18(6):840–8. [78] López-Otín C, Bond JS. Proteases: multifunctional enzymes in life and disease. J Biol Chem. 2008;283(45):30433–7. [79] Imada C, Simidu U, Taga N. Isolation and characterization of marine bacteria producing alkaline protease inhibitor. Nippon Suisan Gakkaishi. 1985;51(5):799–803. [80] Kobayashi T, Imada C, Hiraishi A, et al. Pseudoalteromonas sagamiensis sp. nov., a marine bacterium that produces protease inhibitors. Int J Syst Evol Microbiol. 2003;53(6):1807–11. [81] Imada C, Maeda M, Hara S, Taga N, Simidu U. Purification and characterization of subtilisin inhibitors ‘Marinostatin’ produced by marine Alteromonas sp. J Appl Bacteriol. 1986;60(6):469–76. [82] Taichi M, Yamazaki T, Nishiuchi Y. Role of the backbone conformation at position 7 in the structure and activity of marinostatin, an ester-linked serine protease inhibitor. ChemBioChem. 2012;13(13):1895–8. [83] Takano R, Imada C, Kamei K, Hara S. The reactive site of marinostatin, a proteinase inhibitor from marine Alteromonas sp. B-10-31. J Biochem. 1991;110(6):856–8. [84] Laskowski M, Kato I. Protein inhibitors of proteinases. Ann Rev Biochem. 1980;49:9–626. [85] Bode W, Huber R. Natural protein proteinase inhibitors and their interaction with proteinases. Eur J Biochem. 1992;204(2):433–51. [86] Maynes JT, Cherney MM, Qasim MA, Laskowski M, James MNG. Structure of the subtilisin Carlsberg-OMTKY3 complex reveals two different ovomucoid conformations. Acta Crystallogr Section D Biol Crystallogr. 2005;61(5):580–8. [87] Kanaori K, Kamei K, Taniguchi M, et al. Solution structure of marinostatin, a natural ester-linked protein protease inhibitor. Biochemistry. 2005;44(7):2462–8. [88] Taniguchi M, Kamei K, Kanaori K, et al. Relationship between temporary inhibition and structure of disulfide-linkage analogs of marinostatin, a natural ester-linked protein protease inhibitor. J Peptide Res. 2005;66(2):49–58. [89] Taichi M, Yamazaki T, Kawahara K, et al. Structure-activity relationship of marinostatin, a serine protease inhibitor isolated from a marine organism. J Peptide Sci. 2010;16(7):329–36. [90] Taichi M, Yamazaki T, Kimura T, Nishiuchi Y. Total synthesis of marinostatin, a serine protease inhibitor isolated from the marine bacterium Pseudoallteromonas sagamiensis. Tetrahedron Lett. 2009;50(20):2377–80. [91] Miyamoto K, Tsujibo H, Hikita Y, et al. Cloning and nucleotide sequence of the gene encoding a serine proteinase inhibitor named marinostatin from a marine bacterium, Alteromonas sp. strain B-10-31. Biosci Biotechnol Biochem. 1998;62(12):2446–9. [92] Arnison PG, Bibb MJ, Bierbaum G, et al. Ribosomally synthesized and post-translationally modified peptide natural products: overview and recommendations for a universal nomenclature. Nat Product Rep. 2013;30(1):108–60.

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Eva Nordberg Karlsson, Roya R.R. Sardari, Emanuel Y.C. Ron, Snaedis H. Bjornsdottir, Bjorn T. Adalsteinsson, Olafur H. Fridjonsson and Gudmundur O. Hreggvidsson

3 Metabolic engineering of thermophilic bacteria for production of biotechnologically interesting compounds Abstract: Many thermophilic bacteria are efficient biomass degraders (producing polysaccharide degrading enzymes and utilizing a great variety of substrates, e.g. lignocellulosic polymers, pentoses, hexoses, as well sugar acids, and sugar alcohols). This makes them interesting organisms as potential cell factories in a circular bioeconomy. Lignocellulosic and marine macroalgal biomasses are regarded as sustainable biorefinery feedstocks for the production of energy carriers and platform and specialty chemicals, thereby meeting impending fossil fuel shortage and counteracting accumulation of greenhouse gasses. However, progress in using thermophilic bacteria that utilize these feedstocks as carbon sources has been hampered by the lack of suitable engineering tools to improve the production profiles of interesting target metabolites as specific synthetic production pathways need to be inserted/modified or existing pathways optimized by metabolic engineering. In this chapter, we review the progress on the use of thermophilic bacteria in metabolic engineering and the available engineering tools and give examples of species for which successful engineering has been accomplished. Today, the majority of thermophilic bacteria targeted for production of compounds of industrial interest by metabolic engineering belong to the phylum Firmicutes (e.g. Thermoanaerobacterium, Caldocellulosiruptor, Geobacillus, and Bacillus), taking advantage of anaerobic catabolic pathways producing organic acids and alcohols. However, there are additional and aerobic species gaining interest concerning biomass degradation and the ability of carbon dioxide fixation as well as production of molecules of interest, and some examples of this are also given.

3.1 Introduction The potential of enzymes from thermophilic microorganisms has been highlighted for many years and has also resulted in technological interest due to the possibility to expand the window of operation for enzyme catalysis in industrial biotechnology. As a consequence, numerous enzymes from thermophiles have been characterized after cloning and expression in heterologous mesophilic hosts, and a number of enzymes have also been commercialized for use as biocatalysts [1]. The next step in utilizing

https://doi.org/10.1515/9783110424331-003

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thermophiles has come with the gathering of enormous amounts of genomic and metagenomic sequence data for extreme thermophiles, providing a source of genes encoding additional putative biocatalysts for a wide range of biotransformations under harsher, but perhaps more suitable, conditions than can be used today. So far, the focus been directed toward finding novel enzymes or enzyme groups. When moving toward multistep bioconversions, an intact microbial cell is, however, preferable and usually necessary for economically viable processing. While the potential of thermophilic enzymes has been recognized for a long time, thermophilic bacteria as bioconversion organisms have received less attention. Bioconversions, requiring cultivation and production of metabolites from whole microorganisms, have still mainly targeted mesophilic microorganisms that are easily cultivated and genetically engineered, e.g. the prokaryotes Escherichia coli, Bacillus spp., and the yeasts Saccharomyces cerevisiae and Pichia pastoris [2–5]. Many of these mesophiles are well-proven and efficient hosts for substrates with glucose as the prime carbon source, but they are far less suitable for mixed carbohydrate substrates derived from next-generation feedstocks. Key properties in terms of sugar uptake or metabolism may be missing or inefficient. They may lack extracellular polysaccharide hydrolyzing enzymes and have limited enzyme secretion abilities such as in E. coli. These organisms are also derived from a narrow environmental range regarding temperature and pH, resulting in low tolerance to adverse physicochemical conditions. These established mesophilic organisms are therefore suboptimal for many processes involving carbohydrate substrates in nextgeneration feedstocks.

3.2 Increased interest in thermophilic systems in a circular bioeconomy Geothermal areas are an important source of robust organisms preadapted for harsh and fluctuating conditions encountered in biorefinery processes and feedstocks. Many lineages of thermophilic bacteria are adapted to conditions of extreme pH and to the presence of toxic sulphuric compounds and poisonous metal ions and complexes – conditions that also may be encountered in relatively raw biomass feedstocks fed to bioreactors. Thermophilicity of organisms has potential advantages from a processing perspective, as the ability to grow at high temperatures in bioreactors reduces costs of cooling, distillation, and extraction and prevents contamination of spoilage bacteria. Fermentation at elevated temperatures also eases extraction of many volatile products either by distillation or gas stripping, which alleviates the potential problem of product inhibition or intolerance and should prolong the fermenting life of cultures. Moreover, high temperature increases the solubility of polysaccharides, leads to reduced viscosity of fermentation broths, and facilitates

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enzymatic access to polysaccharides. This subsequently mitigates fermentation scaleup problems of mixing and aeration and enables greater feedstock loads [6]. Despite the number of potential processing advantages listed above, it is the ability of thermophilic bacteria to utilize complex and polymeric lignocellulosic carbohydrate sources that has been a driving force for developing engineering systems for thermophilic bacteria. Many heterotrophic thermophiles are versatile producers of polysaccharide degrading and modifying enzymes and utilize all of the lignocellulosic sugars, pentoses, and hexoses (Tab. 3.1), as well sugar acids and sugar alcohols found in macroalgal marine biomass. The capability of direct utilization of a variety of polysaccharides from biomass thus also qualifies a number of thermophilic bacteria as inherently suitable candidates for one-step consolidated bioconversion processes (CBPs) or simultaneous saccharification and fermentation (SSF) processes [7–9]. SSF and CBP may have advantages over a separate hydrolysis and fermentation process, including maintaining glucose concentration at relatively low levels, which is favorable for many bacteria, avoiding inhibiting degradation products that are a consequence of harsh pretreatment of the biomass, and reduced process investment costs as only one reactor is used instead of separate ones for hydrolysis and fermentation. Albeit the increasing interest, relatively few thermophilic species have yet been engineered to bioconversion organisms. There are several reasons for this: First, there has been a lack of genetic tools and transformation protocols. Second, the available knowledge on metabolic systems in thermophilic bacteria has been limited. Anabolic pathways, product range of catabolism pathway, and the presence and types of secondary metabolite pathways have only been studied in a few key species. This has led to that thermophilic species/strains of mapped abilities, and possibilities for particular production targets are still rather few (with many of the best known candidates found in the phylum Firmicutes, Fig. 3.1a). Third, many organisms need significant development of the cultivation technology to be competitive compared to the traditional workhorse organisms (e.g. E. coli and S. cerevisiae). However, things are progressing and include the development of transformation techniques and genetic tools for important thermophilic bacteria. Examples of engineered species now include heterotrophic anaerobic species belonging to the genera Thermoanaerobacterium [10–12], Thermoanaerobacter [13], Clostridium (renamed Ruminiclostridium) [14], Caldicellulosiruptor [15], and Thermotoga [16] (Fig. 3.1a), all of which produce a diversity of polysaccharide degrading enzymes. Heterotrophic aerobic species include bacteria classified under the genera Thermus [17, 18], Rhodothermus [19–21], and Geobacillus (a facultative bacterium) [22], with the former having natural competence, while the two latter are species with a wide range of polysaccharide degrading enzymes. While most of the target thermophiles have a wide substrate range, the metabolic engineering effort is not limited to product formation. It also includes modifying, improving, and/or extending substrate utilization range to improve efficiency by increasing the reducing power of the catabolic pathway in anaerobes. This has for example been

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demonstrated in Thermoanaerobacter mathranii, by enabling the species to utilize glycerol. Glycerol is a waste material produced in large quantities in biodiesel production, and close to having a negative price. By introducing the gene for glycerol dehydrogenase into the organism, higher ethanol yields were observed from xylose [13]. Recently, the interest in engineering thermophilic bacteria has also gone beyond direct biomass utilization, including engineering of organisms with ability of carbon dioxide fixation, such as the hydrogen-utilizing autotrophic bacterium Hydrogenobacter thermophilus [23] and recently also the heterotroph C. thermocellum [14], opening possibilities for production beyond the current limits of sustainable outtake of biomass. In all cases, new strategies for the development of genetic tools, along with high throughput omics and analyses platforms, paralleled by the generation of metabolic network models [24, 25] by systems biology and synthetic approaches, enable more

DeinococcusThermus

BACTERIA

Aquificae Aquificae

Thermotoga

Aquifi -cales

Thermo -togae

Hydrogenobacter thermophilus

Bacilli Bacill -ales

Thermo -togales

Bacillus methanolicus

Thermotoga maritima

Bacillus smithii (Para)Geobacillus thermoglucosidasius

Deinococci

Bacteroidetes

Firmicutes Clostridia Thermoanaerobacterial es

Rhodothermaeota

Clostridiales

Rhodothermus marinus

Ruminiclostridium thermocellum

Therm -ales Thermus thermophilus

Caldocellulosiruptor bescii Thermoanaerobacterium saccharolyticum Thermoanaerobacter mathranii

Meta bolic engineering Biomass Metabolome

Genetic network

Sugar

L-Lactic acld

NADH

H2

Sugar

L-Lactic acld

Acetyl CoA

NAD*

Transcriptome Gene

Construction of a new pathway

mRNA

Protein/Enzyme

H2

Acetyl CoA

Acetlc acid Ethanol Acetlc acid Butanol

Ethanol

Products

Fig. 3.1: (a) Thermophilic bacterial species subjected to metabolic engineering. Species from five phyla have been targeted with most efforts so far made using species from the phylum Firmicutes. Species classified in a single order are color coded. (b) General considerations and data to be considered for system-based synthetic pathway engineering.

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directed model-based engineering and open new possibilities that are expected to accelerate research and exploitation of thermophiles as bioconversion organisms (Fig. 3.1b). Obtaining an interesting product or product spectrum is of course a main target, and from this perspective, thermophilic bacteria have commonly been exploited for the same range of metabolites as their mesophilic counterparts. Thermophilic bacteria have thus mainly been in focus for the production of biofuels (e.g. ethanol and hydrogen), selected as a general first target area of metabolic engineering (resulting mainly from anaerobic catabolism). Other building blocks for chemicals (organic acids, such as lactate, but also amino acids) have, with time, come more into focus, and in a few cases, other more complex products have been explored (e.g. pigments) (Tab. 3.1).

3.3 Genetic tools and transformation requirements for engineering in thermophiles While metabolic engineering opens up the possibility to insert desired chemical pathways into any microbial host that has the genetic tools available, it does not guarantee that this will result in industrially feasible production from complex biomass feedstock. Other industrial valuable properties are also needed, such as the ability to cope with adverse and fluctuating conditions, and an appropriate substrate utilization range is required. Zeldes and coauthors [48] have stated in their recent review about engineering possibilities in thermophiles that “it is worth remembering that S. cerevisiae, the current workhorse of bio-ethanol production, came to dominate the field because it was already an excellent ethanol producer,” meaning that the most desirable microbial hosts may be those that already do at least parts of the desired processing well from the beginning. However, the prerequisite for using other species than the established workhorses such as E. coli, B. subtilis, and S. cerevisiae is the availability of genetic tools. The importance of establishing robust genetic systems for the evolution of future thermophilic-based bioprocesses was one of the focus points in the review published by Taylor et al. [49]. Despite that this was in 2011, the list of thermophilic bacteria with suitable genetic tools for engineering has not expanded very much (Tab. 3.1 and 3.2). However, this may not be very surprising as the development of more efficient tools and protocols is normally time-consuming, especially in the beginning of the development, and the tools for selected microorganisms have instead been improved.

3.3.1 Competence and transformation Natural competence (or natural ability to take up and incorporate DNA) is a feature described for some thermophilic bacteria and has been a factor that has resulted in

Aerobic/anaerobic (chemolitho-autotropic) Aerobic/anaerobic (heterotrophic) Aerobic (heterotrophic)

75°C

60°C

50–70°C

50–70°C

Caldicellulosiruptor bescii

Geobacillus stearothermophilus NUB3621 Hydrogenobacter thermophilus (Para)geobacillus thermoglucosidasius NCIMB 11955, DL33 Rhodothermus marinus

Anaerobic (heterotrophic)

80°C

70–80°C

Thermus thermophilus

Aerobic (heterotrophic)

Anaerobic (heterotrophic)

45°C

Anaerobic (heterotrophic)

60°C

Thermoanaerobacter ethanolicus Thermotoga maritima

Anaerobic (heterotrophic)

70–75°C

Thermoanaerobacter mathranii Thermoanaerobacterium saccharolyticum

Anaerobic (heterotrophic)

60°C

Aerobic/anaerobic (heterotrophic) Anaerobic (heterotrophic)

(Rumini)clostridium thermocellum

60°C

Aerobic/anaerobic (heterotrophic)

50°C

Bacillus smithii

Aerobic (methylotrophic)

50–55°C

Bacillus methanolicus

Metabolism

Topt

Organism

Organic acids, D-lactate

Amino acids

Metabolic Products

Sugars, peptides, lipids, triglycerides

Wide variety of carbohydrates

Cellulose, starch, hexoses

Cellobiose, hemicelluloses, starch, etc. Cellulose, cellobiose, xylose, and hemicelluloses CO2-fixation ability Carbohydrates except for microcrystalline cellulose Xylan and biomass-derived sugars

CO2-fixation ability (hydrogen oxidizing) Hexose and pentose sugars, oligomers

(Crystallizable proteins)

Acetic acid, H2

Ethanol, acetate, lactate, H2 Ethanol, acetate, lactate, H2  Ethanol, acetate

Ethanol, lactic acid, acetic acid, H2

Carotenoids

Ethanol, lactate, formate, acetate, pyruvate

–

Cellulose,hemicellulose, lignocellulostic Lactate, acetate, H2 plant biomass, C5 and C6 sugars Hexose and pentose sugars, oligomers Ethanol, lactate, formate, acetate, pyruvate

A variety of carbon sources

Mannitol, glucose, methanol

Carbon source for growth

Tab. 3.1: Examples of thermophilic bacteria with versatile carbon source utilization, used for metabolic engineering.

[17, 45–47]

[42–44]

[40, 41]

[38, 39]

[36, 37]

[14, 34, 35]

[19–21]

[22]

[33]

[32]

[29–31]

[28]

[26, 27]

References

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 79

3.3 Genetic tools and transformation requirements for engineering in thermophiles  Tab. 3.2: Examples of engineering and targets for thermophilic bacteria with developed genetic systems. Organism

Selection/ competency and transformation method (efficiency)

Gene targets

Product target

Reference

Bacillus methanolicus

Protoplast formation, electroporation (103–4)

Cadaverine l-lys

[27]

Caldicellulosiruptor bescii

Auxotrophic: Uracil/5FOA/electroporation, after washing in 10% sucrose Chloroamphenicol/ protoplast formation

Insert lysine decarboxylase from E. coli Engineered aspartate pathway Insert adhE from C. thermocellum Deletion of ldh

Geobacillus stearothermophilus NUB3621 Hydrogenobacter thermogenes

(Para)Geobacillus thermoglucosidasius NCIMB 11955 and DL33

(Rumini)Clostridium thermocellum

Rhdothermus marinus (trpB-) Thermoanaerobacter mathranii

Thermoanaerobacter ethanolicus JW200adhE

Kanamycin/calcium chloride treatment and heat shock

Kanamycin/ Electroporation, high osmolarity washing buffer, with sorbitol and mannitol (103 –106) Thiamphenicol/ electroporation (isoniacin addition) (103) Auxotrophic/ electroporation, glycerol treatment (106) Kanamycin/ electroporation after cellobiose wash buffer. isoniacin addition (102–5) Electroporation-, glycin-, and sucrose-induced protoplast formation, subsequent glycerolcontaining buffer washes (101)

Plasmids and expressionsystem 2-Oxoglutarate: ferredoxin oxidoreductase (OGOR) isoenzyme deletion (Kor, For) Engineered ethanol production pathway (deleted and inserted genes) Expression of adhE genes

Gene deletion in carotenoid pathway Expression of glycerol dehydrogenase and knockout lactate dehydrogenase Modification of adhE expression

Ethanol H2 (acetate)

[69, 70]

–

[32]

Modifying CO2-fixation pathway (Kor necessary for anaerobic growth) Ethanol

[33]

[22, 71]

Ethanol

[72]

Removal of pigment production Ethanol

[21]

Ethanol

[40, 73]

[13]

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Tab. 3.2: (continued) Organism

Selection/ competency and transformation method (efficiency)

Gene targets

Product target

Reference

Thermoanaerobacterium saccharolyticum

Kanamycin/ electroporation, autoplast generation (niacin), subsequent sucrose-containing buffer washes (103–4) Kanamycin/ electroporation, 10% glycerol, 0.85 M sucrose (102) (strain RQ7 natural competence) Kanamycin, uracil/5FOA, electroporation (glycerol treatment), natural competency

Engineered ethanol production pathway (deleted and inserted genes) Recombinant expression of cellulase

Ethanol

[39, 74, 75]

Improve substrate utilization

[16,76]

Protein overexpression

[17, 77, 78]

Thermotoga. sp. (maritima, strain RQ7)

Thermus thermophilus

–

early development genetic systems for the genera Thermoanaerobacter, Thermoanaerobacterium and Thermus. Natural competence has in fact been proposed as a feature for adaptation to extreme temperatures [48, 50], but even when it is present, the uptake is normally not very efficient. Natural competence, however, greatly simplifies transformation procedures: simply mix the DNA to be transferred to a dilute culture of the microorganism, followed by plating at selective conditions. Nevertheless, and due to low natural transformation efficiency, most species with natural competence are today transformed using improved protocols utilizing electroporation (Tab. 3.2). Even if transformation procedures can be solved using techniques like electroporation, this may not solve the issue of introducing foreign DNA into the cell. The introduced DNA is consequently often attacked by restriction endonucleases and degraded once it enters the cell. For example, specific methylation is demanded in Caldicellulosiruptor bescii [48]. This has led to search or construction of strains deficient in the defense system against foreign DNA, such as the restriction deficient strain of the aerobe R. marinus that allows genetic modification [21].

3.3.2 Vectors Most commonly, transfer of genetic material is accomplished using plasmids. Naturally occurring plasmids isolated from related (or the exact) thermophilic species have been used to construct vectors for thermophilic bacteria. Desirable characteristics for

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such vectors include the following features: suitable origin of replication (allowing maintenance in the host), suitable selection marker (allowing selection at thermophilic conditions), reasonable transformation efficiency, as well as features allowing cloning (multiple cloning) and expression (promoters functional in thermophilic species), as well as selection in an established cloning host (normally E. coli), creating shuttle vectors between the established host and the thermophilic species [49].

3.3.3 Selection markers Genetic engineering of thermophiles has been hindered by the paucity of selectable markers because of the thermolability of many antibiotics as well as of the products of the most commonly used resistance genes. This thermolability of antibiotic marker proteins has, in many cases, prevented use in genetic engineering of thermophiles. Thermolability is also dependent on other factors, such as pH and presence/ absence of oxygen. For example, several aminoglycosides remain effective following prolonged incubation in anaerobic growth medium at 75°C, while activity is lost at aerobic conditions. At 72°C, pH 7.3, chloramphenicol, streptomycin, penicillin G, tetracycline, and ampicillin lose activity while kanamycin and neomycin remain active. When reducing the pH (72°C, pH 5), only chloramphenicol and streptomycin remain active [17, 18, 51, 52]. Still, antibiotic selection has been successfully applied in some moderate thermophiles [53–55], which subsequently enabled in vivo selection of more thermostable forms [56, 57]. Thermo-adapted variants (e.g. Bleomycin) have in turn been used as selection agents in more extreme thermophiles (Tab. 3.2). Simvastatin is an unusual example of a thermostable antibiotic, but it is unfortunately not applicable to thermophilic bacteria, as it specifically targets archaeal membranes. However, this compound has been important in developing systems for archaeal species, e.g. Thermococcus and Pyrococcus [48]. Due to the thermolability of most antibiotics and the corresponding marker proteins, selection in thermophiles is often also based on complementation of auxotrophy (Tab. 3.2). Metabolic engineering may require several genetic manipulation steps for improving or instituting the production of a target metabolite, and thus, it is important with alternative markers. Success in metabolic engineering has so far largely depended on the availability of multiple functional markers, and the scarcity of thermostable selectable markers has limited the scope of what could be achieved in thermophiles.

3.3.4 Marker recycling The scarcity of markers may limit the scope of what can be achieved in the reconstruction of a cellular network for more effective production of a target metabolite. Marker

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recycling, however, enables reuse of a particular marker in subsequent genetic engineering work. The marker gene is then removed following the gene insertion/deletion step, by a deletion in a second recombination step. Consequently, the insertion/ deletion event becomes “unmarked” and the marker can be used for another genetic manipulation. As a consequence, most changes are chromosomal, due to the difficulty in finding replicating and stable extrachromosomal plasmids.

3.3.5 CRISPR-Cas9 technology Novel opportunities may come with the implementation of thermostable CRISPR-Cas9 (Clustered Regularly Interspaced Short Palindromic Repeats- and CRISPR-associated [Cas] genes), to engineer chromosomal deletions and insertions in thermophilic bacteria. The CRISPR immune systems destroy invading foreign DNA and are guided by specific RNA complexes that recognize foreign DNA [58]. In prokaryotes, these systems can broadly be divided into six different types [59, 60], which differ in the types and number of Cas proteins acting in concert with the guiding RNA complex for recognition and destruction of foreign DNA. Type II systems have only one Cas protein, Cas9, for recognition and destructive cleavage of foreign DNA [61]. This has made the Cas9 enzyme and associated type II components the ideal and highly successful choice for construction of genome editing tools for a variety of mesophilic organisms but has only recently been adapted for efficient use in thermophilic systems. For mesophiles, an efficient genetic engineering tool (SpyCrisprCas9 system) has been constructed using CRISPR-Cas components from Streptococcus pyogenes. It utilizes the type II Cas9 enzyme and a chimeric molecule (sgRNA) for target DNA recognition [62] that is constructed from the crRNA (CRISPR RNA; for target recognition) and the tracrRNA (trans-activating crRNA; for Cas9 recognition). The crRNA includes a short complementary sequence (the spacer) to the site targeted for cleavage (the proto-spacer). The recognition and the following double strand cleavage rely on the presence of two nuclease domains in Cas9, a RuvC-like nuclease domain located at the amino terminus and a HNH-like nuclease domain that resides in the mid-region of the protein [63]. Furthermore, the Cas9 enzyme has a C-terminal domain that recognizes the so-called protospacer adjacent motif (PAM), a 3–8-nucleotide-long sequence [64]. The PAM sequence needs to be present for cleavage to occur in the adjacent protospacer sequence. Maximum functional temperature for the SpyCrisprCas9 system is, however, about 45°C, which has limited its use to mainly mesophilic organisms. Thermostable CRISPR-Cas9 systems have recently been identified and characterized. Homologues to Cas9 (Csn1) have been recognized in the thermophilic genera Geobacillus [65, 66], Acidothermus [67], and Ignavibacterium [68], and their potential for use in genetic engineering has been investigated. This involves the identification of necessary components of the respective immune system, the crRNA, the tracrRNA

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sequence, the required PAM sequences in infecting viruses, and subsequent construction of the Cas9 guiding sgRNA. The coauthors of this chapter from Matis ohf have also recently cloned, expressed, and purified a Cas9 protein from Geobacillus strain LC300. The protein is a thermostable, targetable DNA nuclease. It is currently being adapted for use for genome editing of extreme thermophiles. Similarly, the Cas9 protein from the thermophilic bacterium Geobacillus stearothermophilus catalyzes RNA-guided DNA cleavage up to 70°C in vitro, and by replacing the PAM recognition domain in this protein with the homologous domain from Geobacillus LC300, the PAM recognition specificity was altered [65]. The third thermophilic species used as a basis for the development of a CrisprCas9 system is Geobacillus thermodenitrificans T12, also based on the type II system [66]. This showed optimal activity at 60°C in vitro and detectable activity after preheating for 1–2 minutes at 65°C. Interestingly, a Cas9/sgRNA RNP complex preassembled before activity testing showed good activity at 70°C. The CRISPRCas9 system from G. thermodenitrificans was then adapted for use in the moderate thermophile Bacillus smithii. Knock-out of two genes in B. smithii growing at 55°C was attempted with the system, resulting in complete lethality for one gene and predominantly in clones with a mixed KO/WT genotype for the other gene [66]. In summary, thermophilic CRISPR-Cas9 editing tools based on the type II immune system from the genus Geobacillus are functional at 60 to 70°C, a temperature range suitable for genetic manipulation/engineering of most thermophilic bacteria. As the Cas9 enzyme was shown to be stabilized in the RNP complex, further stabilization may be expected in the environment of the cytoplasm by the presence of compatible solutes and chaperone proteins. Thus, thermostable CRISPR-Cas9 system adapted to thermophilic species, such as Thermus, Rhodothermus and Thermoanaerobacterium, can now be made. This will accelerate progress in metabolic engineering of thermophiles, circumventing the need of a wide array of selective markers or cumbersome and time-consuming marker recycling methods.

3.4 The importance of metabolic range – anaerobic/aerobic systems for different types of products Bacteria undergo different types of reactions to provide energy for growth and other activities. Important for the success of engineering is of course, in all cases, knowledge of the metabolic routes in the respective organism. By a systems biology approach, there are thus today possibilities to establish growth characteristics of thermophilic bacteria, combined with extensive transcriptomic, metabolomic, and proteomic data that serve as aids to generate genome-scale models that guide strain construction (Fig. 3.2). The data from these physiological omics studies can then be used to both improve the genome-scale metabolic model and generate transcriptional regulatory network models. This will establish a rational basis for describing the capacity of the organism that guides the metabolic engineering work.

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Genome sequence

DRAFT RECONSTRUCTION

Annotation

Genetic data Biochemical data

CURATED RECONSTRUCTION

Metabolite data

Omics data

GENOME-SCALE METABOLIC MODEL

DESIGN & DEVELOPMENT

Fig. 3.2: Genome-scale metabolic reconstruction summarized in four major phases, each building the previous.

The selection of chassis organism for metabolic engineering for a particular process should primarily be based on the “metabolic capacity” of the organism, the substrate utilization range required for a particular feedstock, and the (potential) product range of the organism. Anaerobes are therefore usually the organisms of choice for the production of low-value, high-volume commodity chemicals. They produce small organic acids and alcohols that are waste products of catabolism due to incomplete oxidation of substrate and further reductive reactions for regeneration of NAD. Consequently, anaerobes exhibit lower cell yields than aerobes do, but conversely, their end-product yields are generally much higher. During anaerobic metabolism, approximately 95% of the glucose is converted to organic acids and ethanol, while less than 5% is converted to cell mass. The growth rate usually also decreases at anaerobic conditions [79]. Heterotrophic aerobic organisms are capable of complete oxidation of carbohydrates to carbon dioxide, making use of the TCA cycle and the respiratory chain in addition to the glycolytic pathway. They are far more efficient in NAD regeneration and in ATP production and therefore suitable for the production of complex primary and secondary anabolic compounds. High growth rate and conversion of carbohydrates into high levels of cell mass and large amounts of carbon dioxide are characteristic of aerobic metabolism. In aerobes, the ratio of biomass and carbon dioxide is typically 50:50. Due to these characteristics, aerobic microbes may be the organisms of choice for most specialty chemicals, i.e. high-value, low-volume chemicals.

3.4 The importance of metabolic range – anaerobic/aerobic systems 

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To date, most genetic engineering efforts have been made using anaerobic biomass converting candidates from the phylum Firmicutes for production of ethanol (Fig. 3.1, Tab. 3.1 and 3.2). Within this phylum, it has, however, also been possible to combine the high rate of glycolysis and low cell yield of anaerobic fermentative metabolism with the high growth rate and availability of an external electron acceptor of aerobic metabolism for the production of oxidized and redox neutral compounds, which is also important for industrialization. In the anaerobic phase, the production of oxidized compounds is expected to increase product yield and product formation rate concomitantly with less production of carbon dioxide and cell mass. The major advantage for the production of reduced compounds would be increased process stability due to the elimination of the energy advantage of NADH oxidation by the electron transport chain. To address the possibility of combining the beneficial effects of aerobic and anaerobic metabolism into a single biocatalyst, the facultative anaerobic strain Geobacillus thermoglucosidasius has been used for biofuel production [13, 22, 74]. More recently, non-photosynthetic routes for biological carbon dioxide fixation have received increased interest, with the possibility to create “electrofuel”-producing organisms that produce valuable industrial precursors via carbon dioxide fixation [23], and also in this area, there are interesting thermophilic bacterial species. These species include, for example, the autotrophic Hydrogenobacter thermophilus, as well as the heterotrophic cellulose degrader Clostridium thermocellum (Tab. 3.2), both with some engineering tools available. One-carbon (C1) reductive metabolism was very recently reported in C. thermocellum, together with critical enzymes responsible for fixing CO2 capable of channeling fixed carbon to the C1-metabolic pathway, paving the way for engineering of simultaneous cellulose and CO2 utilization (which could improve the theoretical limitation in carbon efficiency) [14]. Below some examples of metabolic engineering efforts in thermophilic bacteria are given, showing in more detail efforts on specific species.

3.4.1 Thermoanaerobacterium spp. for ethanol or 1,2-propandiol production Thermoanaerobacterium is a genus of Gram-positive anaerobic bacteria. Compared to Escherichia coli, Thermoanaerobacterium is a “simpler” organism. It has a significantly smaller genome, and less complex metabolic regulation, shown for instance by co-utilization of pentoses and hexoses. The bacterium grows at a temperature range of 50–72°C and the pH range is 4.5–6.5. Species in this genus show diverse metabolic activities and efficient fermentation capacities and are easily cultivated [10–12, 80–82]. Strains of Thermoanaerobacterium can easily be transformed with host vectors using electroporation and are also naturally competent [11]. Consequently, Thermoanaerobacterium species have been chosen as model organisms for the development of microbial bioconversion systems from second-generation

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biomass feedstock (lignocellulosic biomass) for the production of the bulk and platform chemicals ethanol and 1,2-propanediol (1,2PD). Species of Thermoanaerobacterium have a broad substrate range utilizing a variety of hexoses and pentoses – including glucose, mannose, galactose, xylose, and arabinose, as well as cellobiose [12, 83]. The main fermentative products are ethanol, acetate, and lactate. This means that in the case of ethanol production, the desired product is there in significant amounts from the start. The wild-type strain of Thermoanaerobacterium strain AK17 is a very efficient ethanol producer, reaching 75% of the theoretical yield on glucose under optimal conditions [12]. In previous work, lactate and acetate production was eliminated in T. saccharolyticum and strain AK17 by chromosomal gene knock-outs [10, 74, 84]. The ace- strain produced mostly lactate and the double knock-out mutant (lac-, ace-) only ethanol, and additional reducing power could be obtained by deleting the hydrogenase [75]. Thermoanaerobacterium species also produce 1,2PD from mono-sugars. 1,2-PD (or propylene glycol) is a major commodity chemical used in the synthesis of biodegradable plastics and polymers, with a global demand estimated to be around 1.36 Mt/annum for several industries [85]. The interest in 1,2-PD increases since it is less toxic than products based on ethylene glycol for humans and animals. The US Food and Drug Administration has determined 1,2-PD to be “generally recognized as safe” for use in food, cosmetics, and medicines [85, 86]. The yields of 1,2PD obtained in Thermoanaerobacterium exceed production in strains engineered for 1,2PD in e.g. yeast and E. coli [87, 88]. An additional feature of Thermoanaerobacterium spp. is that it is one of two known bacterial genera that naturally produces enantiomerically pure R-1,2PD. Pure stereoisomers have a considerable additional value as chiral building blocks for the synthesis of specialty chemicals, such as optically active propylene oxide and polymers. R-1,2PD is, for example, an intermediate in the synthesis of chiral pharmaceutical products such as Tenofovir [89, 90], and production of this intermediate can replace chemical synthesis.

3.4.2 Geobacillus thermoglucosidasius, a facultative organism engineered for ethanol production Geobacillus thermoglucosidasius is a Gram positive thermophile reported to exhibit rapid growth and capable of utilizing a variety of plant-derived feedstocks. Geobacillus has been more investigated in recent efforts to engineer more efficient ethanol producers [22, 74]. Ethanol is produced in this species, but less efficient than in S. cerevisiae. In Gram-positive bacteria (of which G. thermoglucosidasius is an example), ethanol production is also accompanied by organic acid production (which could be explored as other putatively interesting products). Both thermophilic anaerobes and anaerobically grown facultative microbes are efficient in ATP

3.4 The importance of metabolic range – anaerobic/aerobic systems 

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generation from glucose, making use of additional pathways that lead to a number of products: ethanol, acetate, lactate, CO2, and H2 [91]. Engineering efforts have been done that channel the pathways toward either of the two products, ethanol or lactate, although the main efforts have thus far been focused on ethanol production. When grown on glucose, acetyl-coenzyme A (CoA) is reduced to acetate, and knock-out of the acetyl-CoA/acetic acid reduction pathway has been shown to significantly increase ethanol production in Geobacillus. Pathway modifications involved elimination of both lactate dehydrogenase (Ldh) and pyruvate formate lyase (Pfl) pathways by disruption of the genes ldh and pflB, respectively, together with the up-regulation of pyruvate dehydrogenase expression (pyruvate dehydrogenase is [unlike the situation in E. coli] active under anaerobic conditions in thermophilic bacilli, although suboptimally expressed) [22].

3.4.3 Thermotoga spp. and Caldicellulosiruptor bescii, with less developed engineering systems Thermotoga spp. and Caldicellulosiruptor spp. are versatile biomass degrading thermophilic microorganisms and, for this purpose, are very interesting candidates for microbial biomass conversion. While Thermotoga is among the most thermophilic bacteria known, a species classified as Caldicellulosiruptor are among the most efficient cellulose degrading thermophiles. Thermotoga maritima is a hyperthermophilic anaerobic bacterium that was first isolated from thermal marine sediments in Vulcano, Italy. This Gram-negative bacterium has an optimal growth temperature of 80°C and has an unusually prominent cell envelope (called the “toga,” hence its name) [92]. T. maritima has been shown to metabolize a variety of carbohydrates including glucose, sucrose, starch, cellulose, and xylan. Furthermore, this organism uses sulfur rather than oxygen as an electron acceptor in the metabolism, thus generating hydrogen sulfide (H2S), rather than water, as a by-product of metabolism. With the ability to produce hydrogen from organic waste products, T. maritima is an important candidate organism for the production of renewable energy. Attempts have been made to develop genetic methods for species of Thermotoga. Anti-metabolite resistant and auxotrophic T. neapolitana strains were isolated [93]. Furthermore, shuttle vectors were constructed by fusing the E. coli vector pBluescript with the cryptic Thermotoga plasmid pRQ7 [94]. The vectors pJY1 and pJY2 contained genes encoding chloroamphenicol and kanamycin resistant proteins, which serve as markers in T. neapolitana and T. maritima, respectively. The vectors were delivered to Thermotoga sphaeroplasts via liposomes. However, stable antibiotic resistant transformants were not obtained, probably due to instabilities of both the vectors and the antibiotics under the growth conditions used. Nevertheless, the vectors were maintained in cells grown in liquid cultures for up to 25 generations, as judged by polymerase chain reaction analysis. Recently, recombinant Thermo-

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toga strains were isolated following transformation with a Thermotoga-E. coli shuttle vector. A heterologous kan-gene was functionally expressed and stably maintained in the host strain [16]. This was an important step in overcoming a barrier to genetic manipulation. Nevertheless, effective tools for genetic manipulation are still lacking, which will be an essential requirement for engineering and studies of Thermotoga biology [43]. Caldicellulosiruptor bescii is the first species in its genus that has been successfully transformed and engineered. C. bescii grows optimally at 65–70°C and utilizes various cellulosic substrates [31, 48, 95]. Thus far, selection in the organism has been based on auxotrophy (Tab. 3.2), and an additional challenge has been the requirement to use methylated constructs (or alternatively a construct strain deficient in restriction endonucleases) [15, 70]. As seen in Tab. 3.2, successful engineering has been made and to date includes ethanol production, improved production of hydrogen [70], and a proof of concept for tungsten assimilation (expressing an archaeal tungsten containing protein) [96]. In 2014, direct conversion of switch-grass to ethanol was demonstrated [69]. Wild-type C. bescii does not produce ethanol, while for an engineered strain, even direct conversion from switch-grass (2%) to ethanol (12.8 mM and 70% of the fermentation products) has been proven. The engineering efforts for this purpose included knock-out of the gene encoding Ldh, combined with insertion and heterologous expression of a bifunctional acetaldehyde/alcohol dehydrogenase from Clostridium thermocellum [69].

3.4.4 Thermus thermophilus and Rhodothermus marinus – natural pigment producers Thermus thermophilus and Rhodothermus marinus are both aerobic bacteria. Their product profile is hence different compared to the above-mentioned thermophilic species, which utilize anaerobic catabolic pathways, but both species have tools and protocols that allow metabolic engineering (Tab. 3.2). T. thermophilus grows at temperatures ranging from 50 to 82°C, optimum pH 7.5–8, and up to 5% NaCl. Despite that T. thermophilus grows on higher concentration of glucose than other Thermus species (unpublished data) and is often isolated from heated compost, it is limited in its carbohydrate utilization range. T. thermophilus has an unusually high natural competence, and transformation is both medium and growth phase independent (the entire population of cells is competent) [97]. Hence, it can be easily transformed, and carbohydrate utilization traits are easily selected for on plates. Vectors include host fosmid systems for functional screening [98], and modified vectors for high-level expression of glycosidases and for thermoadaptation of alpha-galactosidases have been used [99]. Selection marker systems include auxotrophic markers, and several antibiotic markers and marker recy-

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cling systems have been developed based on pyrimidine auxotrophs [100] and counter selectable marker of a conditionally lethal mutant allele (pheS) [18]. An efficient marker recycling system has recently been developed for T. thermophilus HB27 based on markerless gene deletion with cytosine deaminase (codA) [101]. Genetic engineering for functional studies in Thermus has, in most cases, involved insertional mutagenesis [18, 102], and recently, transposon mutagenesis was developed in T. thermophilus [18]. This gives T. thermophilus great potential for engineering, but despite that fact, actual production trials of metabolites are, to our knowledge, not reported. Instead, the developed systems have been used for the production of a variety of proteins [17]. R. marinus is an aerobic heterotrophic thermophilic bacterium isolated from a coastal geothermal area in Iceland and grows at temperatures of up to 77°C. It produces compatible solutes and tolerates at least 7% NaCl [103, 104]. It has been subject of considerable research, much of which has been devoted to its thermostable enzymes on account of their biotechnological potential. R. marinus grows on lignocellulose sugars (except cellulose) and secretes a variety of hemicellulases and amylolytic enzymes. Examples are xylan, glucan, and mannan degrading enzymes, some of which have been studied in great detail [105–111]. R. marinus exhibits reproducible growth on solid defined media, and a restriction deficient isolate has been selected as a recipient for gene transfer experiments [112]. The thermoadapted kanR gene, the selection marker of choice for genetic work on thermophiles is, however, unsuitable for R. marinus because of its natural resistance to aminoglycosides. Instead, selective markers have been developed using trpB and purA encoding enzymes in the tryptophan and adenine biosynthetic pathways, respectively. In engineering experiments, the endogenous trpB and purA genes were deleted from the chromosome (ΔtrpBΔpyrA), making it compatible with both Trp+ and Ade+ selection. A small, cryptic R. marinus plasmid, pRM21, of 2935 bps was used to construct R. marinus-E. coli shuttle vectors, incorporating the R. marinus trpB gene expressed from the promoter of the groES/L heat shock operon, allowing heterologous expression of genes as well as induction by temperature shifts [19, 20, 113]. Inframe deletions using the trpB and the purA marker genes were made, and selection efficiency of the strain was then demonstrated by insertional mutagenesis of the carotenoid biosynthesis gene crtBI, resulting in colorless Trp+, CrtBI- mutants [21]. The mutagenesis thus targeted the carotenoid pathway, which is of increasing industrial interest due to its pigments. Both R. marinus and T. thermophilus produce carotenoids via lycopene using the terpenoid pathway including γ-carotenoid acyl glucoside and a yellow β-carotenoid acyl glucoside [114, 115]. Together, these organisms constitute a gene pool encoding complementary enzymes for producing a variety of terpenoids and carotenoid derivatives of industrial value such as echinenone, cantaxanthin, zeaxanthin, and astaxanthin, e.g. as supplements in feed and food [116].

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3.5 Future perspectives Thus far, the most advanced efforts on metabolic engineering of thermophiles have been focused on microbial production of biofuels using renewable feed-stocks. However, economical production of a low-value bulk compound, e.g. ethanol, is today challenging. More diverse product-portfolios including higher-value compounds hence receive increased attention as targets of microbial productions. Such approaches will, however, require careful analysis of both the possibilities and limitations for production of desired metabolites in targeted host organisms. To increase production efficiency, optimization of both metabolic efficiency (i.e. efficient carbon flux toward product, combined with a high oxidation of the substrate (generating ATP and NAD(P)H) will be advantageous. This will most likely result in the development of different host organisms for different types of products, and in this range, we believe that thermophilic bacterial production systems will be selected for some future processes. To stimulate the development of efficient thermophilic bacterial production organisms, systems analysis of selected candidates is important (covering aspects ranging from the genetic scale to the fermentation engineering), allowing the above-mentioned optimization of metabolic efficiency, and a broad metabolic knowledge is foreseen to gain a natural role in the selection and development of promising thermophilic cell factories. Acknowledgments: Financial support from the ERA-net project Thermofactories and the EU-projects Macrocascade and VirusX is gratefully acknowledged. GOH and OF also acknowledges the support from the EU-project MacroFuel and ENK acknowledges support from the Swedish Research Council Formas and the Novo Nordisk Foundation.

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Giannina Espina, Paulina Cáceres-Moreno, Daniela Correa-Llantén, Felipe Sarmiento and Jenny M. Blamey

4 Extremozymes: from discovery to novel bio-products 4.1 Abstract Biocatalysis has proven to be an essential industrial tool for converting raw material into valuable bio-products. The discovery of new enzymes, the improvement of enzymatic features, and the development of new processes for enzyme production will drive future innovation. For the chemical industry, the use of enzymes presents important benefits, which include higher selectivity, increased sustainability, and a low toxicity. These benefits are translated in cleaner production processes and lower environmental impact. Enzymes derived from extremophiles, or extremozymes, often have extraordinary properties, which include being able to carry out reactions at nonstandard conditions (e.g. high or low temperatures, acidic or alkaline pH, high concentrations of salt or organic solvents, and high pressure) where other enzymes underperform. Working with extremophiles and their native extremozymes is difficult due to the culture conditions required; they have usually low cell yield and low enzyme expression. For these reasons, to achieve industrial production levels, available extremozymes have been overexpressed in suitable heterologous host-vector systems. In this review, we present a road map to find enzymes from extremophiles. We will explore the process from discovering an enzymatic activity in a microbial crude extract through the application of a functional biochemical approach, up to the development of a new enzymatic product.

4.2 Biocatalysis benefits and barriers Enzymes are biocatalysts that improve the rate of biochemical reactions by lowering their required activation energy. A reaction that would normally take a long time, such as oxidation, can occur in just milliseconds if the appropriate enzyme is used. Due to the high selectivity of enzymes, it is possible to optimize chemical reactions in different manners: maximizing the use of energy, minimizing the generation of secondary products, and reducing the reaction time. Thanks to the commercial, technical, and environmental benefits that the use of enzymes offers, biocatalysis is increasingly gaining interest for application in several industrial processes. https://doi.org/10.1515/9783110424331-004

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Compared to conventional chemical processes, the implementation of biocatalysis in mature and well-developed existing industrial processes allows reduction in operating costs and increases efficiency in manufacturing processes, therefore creating better profitability. Moreover, the use of biocatalytic processing routes offers additional benefits for the environment, as well as the safety of the staff by avoiding the utilization and handling of many toxic compounds and solvents, normally employed in chemical processes (e.g. dichloromethane, dimethylaniline, chloroacetyl chloride, phosphorous pentachloride, and phosphorous trichloride). For example, comparing the efficiency and environmental aspects to produce the antibiotic cephalosporin (7-aminocephalosporic acid [7-ACA]) chemically and enzymatically, the first process showed a higher yield (51 w/w%) than the second (46 w/w%) but a lower reaction mass efficiency and half mass productivity (1.2% compared to 2.3%, excluding water). Furthermore, the chemical process also uses more hazardous materials, approximately 60% more energy, 40% more oil equivalents, 16% more mass, double the greenhouse gas, impact and about 30% higher photochemical ozone creation potential and acidification level than the enzymatic one, thus generating a greater environmental impact [1]. Currently, the change from chemical catalysts to the use of enzymes is driven by regulations and incentives from the governments of different countries encouraging the generation of green industrial processes using bio-products. As an example, the US Environmental Protection Agency, the US Council on Environmental Quality, and the US Department of the Interior are developing and creating regulations to reinforce industrial environmental responsibilities. To date, one of the major barriers to enter in the industrial enzymes market is the effort required to sustain the presence of a biotechnology company in a global and dynamic market. In addition, even though enzyme technologies have improved over the last decades, there is an important need to generate more molecular tools and keep refining enzymatic processes for different applications. The understanding of specific customer needs in terms of operational ranges, efficiency, and bioprocessing is critical for success. Also, not all companies are willing to test novel enzymes that can improve their processes, if this means a large change in their current activities. Sometimes stopping a process for just a few days, so that a novel biocatalytic step can be implemented, is extremely expensive. However, the risk to incorporate a novel process may be compensated by additional benefits including social and marketing positioning by being recognized as a “green” company when reducing their environmental impact [2]. Also, an important reduction in production costs could be achieved in the long run by implementing enzymatic steps or full biological processes that use less energy and water or generate less waste. Additionally, commercialization strategies, as well as an efficient management, are among the most critical parameters to be competent and resourceful in marketing novel bio-products. Another challenge is to achieve large-scale production under the quality required by the specific market. Understanding the volume of

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production and the quality of an enzyme is a case-by-case situation, which depends on the application of interest. Any commercialization strategy must take into consideration different scenarios, which may include market size, pricing structures, production formats, and environmental and safety regulations, among others. A project evaluation considering all the above-mentioned benefits and drawbacks, as well as constant market intelligence, is required to develop new processes and utilize new technologies using biocatalysts.

4.3 Extremozymes discovery and development functional roadmap Extremophiles are microorganisms that thrive in environments that present extreme physicochemical conditions such as high and low temperatures, acidic or alkaline pH, salinity, high pressure, high radiation, high concentration of metals, among others. In order to adapt to these environments, extremophiles have developed special mechanisms, including novel biocatalysts and unique metabolic pathways, that allow them to respond to harsh conditions [3]. Most of the currently available commercial enzymes are of mesophilic origin. They present optimal activity in narrow ranges of conditions, which limits their actual application under industrial settings. Enzymes from extremophiles, also known as extremozymes, offer an efficient alternative to overcome several barriers present in industrial biocatalysis. For example, extremozymes are more stable and robust than their mesophilic counterparts, allowing them to perform catalysis under the harsh conditions found in most industrial applications. In order to find novel microorganisms with interesting enzymes for industrial applications and to generate novel biocatalysts, the two most common approaches are metagenomic screening and direct exploration of enzymatic activities [4]. Both approaches involve environmental sampling as a starting point, which provides a snapshot of the diversity and density of microorganisms from a particular location, in a determined space and time. Environmental samples taken from places with extreme environments, like glaciers, geysers, and hydrothermal vents, among others, provide better chances to find microorganisms adapted to specific extreme conditions that could resemble conditions of industrial interest. Sampling extreme environments might be challenging due to the difficult access to interesting places. Eventually, protected places require special licenses in order to take samples and to work with them (e.g. Antarctic, natural reserves). After obtaining suitable samples, it is possible to follow a metagenomic route that comprises the genomic analysis of the microorganisms present in the sample by direct extraction, cloning, and sequencing of their DNA without need of culturing, therefore allowing obtaining information from unculturable microorganisms. Metagenomic screening relies on the fact that only a small percentage of the microorga-

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nisms present in the planet have been isolated and studied, and no more than 1% of them can be cultivated [5, 6]. This number is even lower when referred to extremophilic environments. However, it is not possible through genomic methodologies to determine the specific biochemical properties of an enzyme such as specific activity, optimal temperature, pH, and thermal stability [7]. Metagenomic approaches have been extensively reviewed [8–10], and this subject is out of the scope of this chapter. On the other hand, direct exploration of enzymatic activities or functional approach, involves the detection of an enzymatic activity measured on a crude extract of a culturable microorganism. This method, of course, requires the isolation of the respective microorganism beforehand, but the measurement of the enzymatic activity can immediately determine the existence of a bio-catalytic transformation for a given experimental setting, which is an applicable advantage to speed up product development. A strategy for the development of extremozymes is following four phases in a stepwise approach (Fig. 4.1). This includes an initial Discovery Phase, followed by a Development Phase, a Scale-up Phase, and finally a Production Phase. The strategy further detailed in this chapter has provided tangible results for the development of several enzymes, including Swissaustral Microbial Catalase, Microbial Glutamate Dehydrogenase, and Microbial Laccase, as risks can be reduced dividing the process into various steps, offering enough flexibility to decide at the end of each phase the further development and direction of the project to ensure good results before investment is made in the next step.

I. Discovery Phase

II. Development Phase

III. Scale Up Phase

IV. Production Phase

– Isolation of microorganisms from environmental samples. – Identification of microorganisms (16S and genome sequencing). – Development of screening methods/functional approach.

– Purification, activity measurements and biochemical characterization of the native enzyme. – Bioinformatics and identification of enzyme encoding gene. – Development of recombinant extremozymes. – Purification and characterization of the recombinant enzyme.

– Optimization of the biomass and protein yield. – Scale-up of the biomass and protein production. – Scale-up of the enzyme purification.

– Quality controlled biomass generation and enzyme production. – Packaging and labeling of the enzyme. – Final product information (MSDS, datasheet, among others).

Fig. 4.1: Diagram of a stepwise strategy to find novel bio-products.

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4.4 Direct exploration of enzymatic activities 4.4.1 Phase 1 – discovery Isolation of microorganisms from environmental samples When aiming to culture microorganisms, the sampling process must guarantee that the viability of microorganisms and enzymatic activity remains without alteration. Each sample taken from an extreme environment is unique and its processing could influence the growth of different microorganisms. Therefore, to maintain the viability of extremophilic microorganisms and avoid contamination, sampling from soil, water, ice and sediment must not be collected from the surface. The sample must be taken at least at 10–20 cm from the surface, and the use of sterile material and tools is crucial. It is also very important to register the sampling conditions of temperature, pH, and geographical location. In situ processing of the environmental samples increases the possibilities to find unique microorganisms with interesting characteristics. This technique refers to immediately inoculate the environmental sample in culture media, allowing the preservation of a higher number of microorganisms. Depending on the nature of the sample and its original environmental conditions, one or more suitable culture media needs to be selected. Generally, culture media for hyperthermophiles and thermophiles are composed of a high percentage of minerals and low amount of carbon sources. Some of them use hydrogen, thiosulfate, and/or sulfur as electron donor for reducing oxygen and transfer energy to cellular chemical products, as for example, Pyrococcus furiosus medium [11] numbered 377 by the German Collection of Microorganisms and Cell Cultures, DSMZ (Deutsche Sammlung von Mikroorganismen und Zellkulturen) and 1915 by the American Type Culture Collection, ATCC. On the other hand, to grow psychrophilic microorganisms, the required amount of minerals and carbon sources is very low. A useful culture medium specially developed for microorganisms from soils poor in nutrients is the Antarctic Bacterial Medium [12]. In the case of alkaliphilic microorganisms, simple composition media containing Na2CO3 at pH 10 is recommended [13], which allows finding new microorganisms that grow optimally at pH 10 but not at neutral pH. However, for acidophiles, the culture medium needed should contain salts and an acid, such as sulfuric acid, for pH fixation [14]. Cultures for halophilic microorganisms need media low in nutrients but high in salt concentration (i.e. 1.5 M NaCl) and long times of incubation. By varying salt concentration in the media, it has been possible to obtain different halophiles [15]. Culturing extremophilic anaerobes is more complex since their growth often requires mimicking the conditions present in terrestrial volcanic sites or submarine hydrothermal systems, including submarine volcanoes, fumaroles, and oil reservoirs, among others. Volcanic environments usually harbor large amounts of steam, carbon dioxide, hydrogen sulfide, and sulfur and variable quantities

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of carbon monoxide, hydrogen, methane, nitrogen, and traces of ammonia. Strict anaerobes die or immediately stop growing upon exposure to low levels of oxygen. It is therefore important to maintain anoxic conditions during all handling steps of these microorganisms. Most strict anaerobes require not only the absence of oxygen to initiate growth but also a redox potential below −300 mV, which can be only achieved by supplementing the media with reducing agents such as cysteine × HCl, Na2S and rezasurin, a redox sensitive dye that is usually included in the media for culturing anaerobes to monitor the redox potential. Rezasurin is generally nontoxic to bacteria and archaea and is effective at very low concentrations of 0.5 to 1 mg/L [16]. At the laboratory level, it is important to provide media that contain the adequate source of carbon and nitrogen along with trace elements that allow the construction of the metabolic machinery of the microorganism. To avoid oxygen in the media, during the growth, a constant flow of nitrogen or other gases such as CO2 and H2 is necessary. Growth temperatures can range from 60°C to 100°C. It is important to remember that among anaerobes, anaerobic thermophilic and hyperthermophilic archaea are the most extreme anaerobic organisms. If the anaerobes come from a marine environment, a medium that emulates the composition of sea water will be a suitable option. For terrestrial anaerobes, knowing the geochemical composition of the environmental sample to design an adequate medium is desirable. Additionally, microorganisms often live in symbiotic conditions with others, leading to mixed cultures or consortia, for which it is very difficult, if not impossible, to emulate lab conditions that allow them to grow as isolated microorganisms. To overcome these situations, it is necessary to develop new techniques and special culture media. So far, several investigations have been developed in order to resolve these problems, obtaining promising results [17, 18]. I dentification of microorganisms (16S rDNA and whole-genome sequencing) Successfully isolated extremophilic microorganisms need to be properly identified. The identification of new microorganisms is an ambitious work, and a polyphasic approach is needed, including not only morphological, physiological, and biochemical characterization but also the complete 16S rDNA gene sequencing, and its comparative analysis by phylogenetic trees, among others (such as DNA-DNA hybridization studies with related organisms, analyses of molecular markers, and signature pattern(s) and even genome sequencing information and comparison) [19]. In order to obtain the complete 16S rDNA gene sequencing, it is necessary to (i) lyse the microorganism, (ii) extract its DNA, and (iii) perform a polymerase chain reaction (PCR) using appropriate primers. When working with well-known microorganisms such as Escherichia coli or Bacillus subtilis, these are routinely and easily done experiments. However, in the case of many extremophiles, there are several difficulties that make this process nontrivial.

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First, cellular disruption represents a challenge when working with extremophilic microorganisms. For example, microorganisms belonging to the Archaea domain have a unique composition in their lipid membrane, as they have ether links that join glycerol and isoprenoids (hydrophobic lateral chains) [20]. These structures improve the stabilization of these types of microorganisms at high temperatures, because lipid monolayers are more resistant to degradation [21]. Many archaea require not only physical methods (e.g. French press, sonication), but it is also necessary to consider chemical methods like the use of more stringent conditions and lytic buffers containing a mixture of detergents and chaotropic agents. Several protocols have been designed for a wide range of hyperthermophilic microorganisms, which are usually very harsh. In 2013, Mirmohammadsadeghi and collaborators compared five methods to disrupt the hyperthermophilic microorganism Pyrococcus furiosus, showing that the best methods are the ones that combine enzymatic lysis (using proteinase K) and chemical lysis (using SDS) [22]. More intense mechanical methods (e.g. sonication) used to break down the microorganisms could also affect the genetic material, which might generate a DNA smear. On the other hand, psychrophilic and halophilic microorganisms are easier to disrupt. The process can be done with standard protocols adding supplementary buffer washes to remove exopolysaccharides present in the cells. Second, many of the chemicals and conditions required by extremophiles to grow and thrive (i.e. large amounts of salts, metals, sulfur, extreme pHs along with the presence of humic acids) greatly interfere and difficult DNA recovery, which significantly affects the yields of DNA in good conditions that can be obtained. As a consequence, downstream processing such as PCR amplification and sequencing, which heavily relies on the quality, concentration, and purity of the starting DNA material, is also affected. Further purification of DNA extracted from extreme environments is frequently mandatory. Third, even though specific archaeal primers have been developed [23], the majority of universal primers used to amplify the 16S rDNA gene have been designed for bacteria; thus, a high number of members from the archaea domain remain as unknown microorganisms. Besides the sequencing of the 16S rDNA, advances in DNA sequencing technologies from first to second and third generation [24] have allowed obtaining whole-genome sequences of a microorganism in a quick and economical manner. Furthermore, the time and price of whole-genome sequencing are expected to decrease every year, making it more accessible and affordable [9, 24]. Development of screening methods To find the right enzymatic activity, we rely on the technical capability to efficiently assess the biochemical reaction that carries out the transformation of a particular substrate into the desired product. However, to find the right enzyme capable to catalyze, under industrial conditions, a specific transformation of industrial interest,

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a large number of microorganisms/crude extracts or samples need to be screened, and new robust enzymatic assays should be developed every time a novel enzymatic activity is sought. In order for this approach to become industrially useful, setting it up in a miniaturized manner is needed to allow massive and rapid screening and discovery of the most efficient biocatalyst for a specific application. For extremophilic enzymes, miniaturizing the enzymatic tests is not always possible, particularly if we consider the extreme conditions at which the reactions should take place. Materials that are used for these types of methodologies, in many cases, cannot stand the conditions at which extremophilic enzymes optimally work.

4.4.2 Phase 2 – development Activity measurements, purification and biochemical characterization of the native enzyme Activity measurement is the first step to find the right enzyme. Enzyme assays measure either the disappearance of the substrate, or the appearance of the product over time. The majority of the assays currently used are spectrophotometric. But when changes in substrates or products are not observable by spectrophotometric methods because they do not absorb light, coupling assays are used. In this case, light absorbing nonphysiological substrates or products are synthesized. These reactions can be measured by coupling them to enzymes that can be detected via a spectrophotometer. Other optical methods may be used, such as fluorimetry, which is about hundred times more sensitive than absorbance measurements [25]. But only a few enzymatic substrates or products emit fluorescence, such as NADH and some artificial substrate analogues. Another alternative is to assess the product of an enzyme reaction by electro/ radiochemistry or using separation techniques such as high-performance liquid chromatography, gas chromatography, and capillary electrophoresis, depending on their nature [26]. However, these require special and sophisticated equipment and manipulation, increasing the cost of the experiments; for these reasons, they are used when is not possible to apply spectrophotometric or fluorometric assays. For the development of enzyme assays, it must be considered that enzymatic reactions depend on more factors than pH, temperature, and ionic strength. Highly important are the actual concentrations of all the components present in the assay. Since the formation of product is directly connected with the disappearance of substrate, its decline is an adequate measure of the reaction. When two or more products are formed, the determination of only one of the components formed is sufficient to estimate the enzymatic activity [27]. In the case of thermophilic and hyperthermophilic enzymes, it is important to check the stability of the components present in the assay mixture at the temperature required to run the assay [28]. The assay should be operating in a range where the

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initial rate of the reaction is measured and it is proportional to the amount of enzyme added to the assay [27]. Purification of an enzyme requires the design of an experimental procedure which in general involves several steps. The first one is the preparation of the crude extract for which is crucial to establish an optimal cell lysis method. Among the most widely used methods for cell disruption to prepare crude extract are the use of alkali/ heat treatment, French press, and sonication [29]. Once the crude extract is prepared, the purification procedure for enzymes usually involves the use of liquid chromatographic techniques. The implementation of ionic exchange chromatography, affinity chromatography, and size exclusion chromatography is common [30]. An adequate buffer that may contain stabilizers such as glycerol, sucrose, or other sugars is sometimes used along the purification. In the case of anaerobic enzymes, it is very important to carry out all the purification under anaerobic conditions [31]. This involves the use of anaerobic buffers and all the reagents previously degassed, as well as anaerobic chambers containing nitrogen or inert gas atmosphere, maintaining the atmosphere devoid of oxygen. The use of reducing agents in all buffers along the purification is required in many cases [31]. For biochemical characterization, the first parameters to be determined are optimal temperature and optimal pH. Then, it is important to include studies of thermostability, inhibitors, and determination of the kinetic parameters Km, Vmax, and kcat [27, 32]. In addition, determination of stereoselectivity/stereospecificity of an enzyme by using enantiomeric substrates, is also important for industrial context. Furthermore, long time storage is an interesting logistic requirement in industry; therefore, once the enzyme is purified, studies of stability over time at different temperatures for the enzyme in both, liquid and solid/lyophilized form, are needed [32]. Bioinformatics and identification of enzyme encoding genes Bioinformatics have played and will continue playing a significant role in searching of industrial biocatalysts. This can be implemented by DNA and protein sequence search, comparison, and analyses using information present in databases employing different algorithms and software. Even though obtaining a whole-genome sequence is now relatively easy and cost-effective, the crucial work is the bioinformatics analysis of the data [33, 34]. The genome must be assembled and genes need to be properly annotated (often manually curated) in order to identify the gene encoding the enzyme of interest. Genome mining consists in finding the right ORF coding for putative enzymes, through the alignment and comparison with annotated sequences that are deposited in the databases. The fact of trusting in previously annotated sequences brings uncertainty to the expected results, and it might create a bias and hinder the discovery of truly new enzymes [35].

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Development of recombinant extremozymes Industrial processes demand the production of enzymes in large quantities. Currently, using extremophiles as production-strains for novel extremozymes on a large scale is still not feasible due to two main limitations. First, most extremophiles are simply not suitable for large-scale fermentation using bioreactors, especially hyperthermophilic and thermophilic microorganisms, because the biomass that is possible to obtain is, in the majority of cases, less than 1 g/L. This presumably occurs by the accumulation of toxic compounds as a result of Maillard reactions [36] or gases such as CO2 and hydrogen sulfide generated in the presence of elemental sulfur. This translates into obtaining even lower protein yields. Second, operating bioreactors under the extreme conditions of temperature, pH, or salt concentration, necessary for the growth of extremophiles, will shorten the lifetime of sensors and seals, and the high energy and special media required will increase the cost of the process. All of these are counterproductive for the development and production of new industrial enzymes. For these reasons, it has been necessary to use a different approach to obtain a satisfactory yield of an extremozyme of interest. The advances in recombinant DNA techniques opened a new era for protein production; it has been described that 90% of the enzymes currently used on industry (mostly from mesophilic origin) correspond to recombinant versions [4]. Therefore, the current strategy is to clone and express genes derived from extremophiles into well-known heterologous mesophilic hosts. The extremophilic gene to be heterologously expressed must be PCR-amplified or synthetized, taking into consideration the phylogenetic distance between the parental microorganism and the selected heterologous host. Due to the differences that can be found in the gene expression machineries among different taxonomic groups, differences in their codon usage, recognition of promoters, and missing initiation factors and/or cofactors may also affect dramatically the recombinant expression of a functional extremozyme [37]. Special algorithms for codon harmonization are used fairly nowadays, for the synonymous replacement of codons having usage frequencies in the heterologous expression host that are less than, or equal to the usage frequencies of native codons in the native host [38]. To date, the expression of recombinant proteins in Escherichia coli is, by far, the easiest, quickest, and cheapest method. Its physiology, metabolism, and genetics have been extensively characterized, therefore, rapid growth with high cell density can be easily achieved using inexpensive media. Furthermore, there are many genetic tools readily available for its manipulation, with many commercial and noncommercial mutant host strains and vectors, which are optimized for special applications, by means of experimental and computational approaches [39]. Nonetheless, there are other bacterial heterologous hosts available, such as Bacillus subtilis and Bacillus licheniformis, and yeast/fungal systems, such as Saccharomyces cerevisiae, Pichia pastoris, Aspergillus niger, and Trichoderma ressei. These hosts are more commonly used for the recombinant expression of industrial enzymes that require a Generally Regarded as Safe (GRAS) clearance certificate for their applications, or when the

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recombinant expression in E. coli is not recommended (i.e. when a posttranslational modification is needed). For the selection of an adequate recombinant host, the advantages and disadvantages offered by each of the currently available systems must be considered; some of these have been previously reviewed by Gomes et al. in 2016 [40]. Also, it is necessary to choose the appropriate expression tools. Plasmid-based expression is the most popular choice, as genetic manipulation of plasmids is well known, quick, and easy and there are many commercial systems available. In order to select an appropriate plasmid to clone a target gene, the most relevant features to consider are the promoter and the selection marker. A promoter is a specific region of DNA that is located upstream a gene, and initiate its transcription into RNA. Promoters can be constitutively active, which means they are continually turned on, or be more carefully regulated; such is the case of repressible or inducible promoters [41]. These ones are more tightly controlled and the transcription of the target gene can be turned off/on with a chemical agent, heat, or even light. Selection markers are specific plasmid genes that confer a competitive advantage over plasmid free cells and benefit the plasmids stability [41]. The most common ones are genes that provide resistance to a certain antibiotic (e.g. kanamycin, ampicillin, chloramphenicol), which are then added to the culture medium in order to apply a selective pressure that allows discriminating plasmid containing cells [42]. Furthermore, the use of antibiotics also prevents contamination of the culture with other microorganisms. It is important to consider that for industrial production of the target enzyme, the addition of chemicals such as IPTG when using inducible T7 or T5 promoters can increase the cost of the process. An alternative to its use is to employ a culture media specially developed for auto-induction [43]. This media uses lactose and other sugars to induce the expression of genes under the control of T7 or T5 promoters. On the other hand, adding antibiotics to the media increases the costs of the production process, as they are quite expensive and can be degraded or inactivated. Most important, however, is that their presence is undesirable in therapeutic and food products, as well as in the wastewater effluents of the fermentation operation, due to the risk of contributing to antibiotic resistance [42]. Therefore, nowadays, antibiotics-free plasmid systems have been increasingly developed based on the plasmid addiction concept [42, 44]. For the above-mentioned reasons, chromosomal integration is the strategy of choice for the commercial expression of recombinant proteins. Even though integration of a foreign gene in the chromosome is labor-intensive, time-consuming and typically results in lower production rates than with plasmid-based systems (due to a low copy number of the recombinant gene). Resources invested in host development are easily compensated with getting a stable host, able to grow in the absence of antibiotics without any reduction of recombinant protein yields. Furthermore, this approach also had the advantage of not infringing patents [44, 45].

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The expression of recombinant enzymes, especially the ones from extremophilic microorganisms, is not always straightforward and frequently faces additional challenges such as protein degradation, protein toxicity, and protein misfolding [46]. All of these issues noticeably interfere with the production of functional extremozymes, which also argues in favor of the use of a functional approach, as the number of genes to be cloned is drastically reduced to the ones with proven promissory enzyme characteristics. Nowadays, it seems very difficult to anticipate any of the above-mentioned difficulties beforehand, as each challenge may vary from gene to gene and depends on the expression system and host used, which is critical for the successful expression of the gene of interest. For example, in the case of proteolysis, this can be overcome by producing a recombinant protein in protease-deficient heterologous hosts, and in the case of toxic proteins, there are also special hosts available with demonstrated high tolerance to the expression of toxic gene products [47]. The expression of heterologous proteins often reaches nonphysiological concentrations, which may saturate the cellular folding machinery, preventing the polypeptide chain to fold into the correct native three-dimensional structure. This leads to improper secretion in the case of genes containing signal peptide sequences for extracellular expression and/or subsequent protein aggregation and accumulation of misfolded polypeptides within large intracellular aggregates known as inclusion bodies [48]. However, protein aggregation has certain benefits: first, proteins can be protected from proteolysis, and second, it can protect the host by preventing the deleterious effect of toxic recombinant proteins [49]. Also, even though inactive, the proteins inside insoluble inclusion bodies are found to be relatively pure in comparison to proteins expressed in soluble form (either in the cytosol or supernatant). It has been described that recombinant proteins can be successfully recovered from inclusion bodies in active form. The process to ensure the solubilization of the aggregates is performed through a cycle of denaturant and/or reducing agents (e.g. urea, guanidine-HCl, beta-mercaptoethanol, or dithiothreitol), followed by the slow removal of the denaturant agent under conditions that favor refolding and renaturation (e.g. a stepwise process at low temperature, combined with a mixture of reduced and oxidized thiol compound) [50]. However, many times refolding of insoluble proteins results in a very inefficient and expensive process. Furthermore, it is impossible to predict whether a protein will aggregate or not when using a particular host/expression system or how easily it will be solubilized and renatured [45]. Several experimental approaches can be used in order to minimize protein aggregation and to obtain direct expressions of properly folded proteins in soluble and active form. The first one consists of lowering the growth temperature at the mesophilic host’s physiological limit. Another possibility is to change the fermentation medium. Other attempts have been made to produce recombinant proteins in a soluble form

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by co-expression of chaperones and foldases (e.g. Cpn10, Cpn60, GroEL, GroES, thiol-disulphide-isomerases [PDI], and peptidylprolyl-isomerases) that assist with the correct protein folding. Another approach is to express the recombinant protein as a fusion protein with a highly soluble partner (e.g. maltose-binding protein, gluthatione S-transferase, and thioredoxin) [51]. When all the above-mentioned methods fail to increase expression levels, switching to another host/expression system should be considered. Nevertheless, there is a clear need for understanding and developing novel easy-to-use hosts and expression systems in order to be able to express extremophilic enzymes, especially for the expression of phylogenetically distant groups (e.g. Archaea). Kishishita et al. reported in 2015 the use of the fungus Talaromyces cellulolyticus as a host for the expression of hyperthermophilic cellulases from the archaeal Pyrococcus sp. [52]. Some authors have also suggested that new expression systems will have to be developed with extremophilic organisms as the host [53, 54]. Functional systems have been obtained, for example, using the hyperthermophilic archaeon Sulfolobus solfataricus [55]. Genetic improvements of extremophilic expression systems continue to be under intense investigation. Nonetheless, as previously mentioned, in order to use extremophiles as host microorganisms, the development of novel culture media, molecular tools, and engineering technologies is, as well as the design of specialized equipment capable of dealing with extreme conditions.  urification and characterization of the recombinant enzyme P at laboratory scale After obtaining a functional recombinant extremozyme in soluble form, the next step is to purify and characterize it. For this, the preliminary information given by the purification and characterization of the native enzyme, done in phase, has been proven to be very useful. In some cases, the recombinant protein purification can be facilitated by the addition (and subsequent removal) of affinity tags (e.g. polyhistidine tag), which allows reducing the chromatography steps needed, as well as the time and labor involved [56].

4.4.3 Phase 3 – scale-up Optimization of the biomass and protein yield The expression of recombinant extremozymes in easily culturable hosts allows higher productivity in shorter periods of time, reducing production costs. The versatility and scaling-up possibilities for the production of recombinant enzymes open new commercial opportunities for the use of enzymes obtained from extremophiles.

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After obtaining a heterologous host able to express a functional extremozyme in soluble form (either intracellular or extracellular), and having generated its appropriate frozen glycerol stocks as backup, it is necessary to proceed with the optimization of the recombinant expression at laboratory scale. This process is done in flasks with culture volume ranging from 50 mL to 1 L and in small bioreactors up to 5 L. There are many parameters that need to be optimized in order to ensure satisfactory levels of biomass and protein expression, such as culture media, pH, growth temperature, agitation rate, inoculum volume, optimum fermentation volume, culture time, antibiotic concentration (if required) as well as inductor concentration, time, and temperature of induction (if needed). All these parameters are important because, individually and synergistically, they eventually affect the growth, productivity, and production of the bio-product of interest. Conventional optimization approaches involve the study of varying one factor at a time, while maintaining the other factors constant. However, this methodology has shown to be very slow and expensive. For this reason, statistical methods have been developed in order to achieve the optimum value of all the studied parameters and their interactions. The most used statistical method corresponds to the response surface methodology, reviewed by Myers and collaborators in 2016 [57]. This statistical tool allows optimizing several factors at the same time, with minimum material expenses and time. For example, in 2019, Suberu and collaborators improved protease production in Bacillus. They optimized different factors such as carbon and nitrogen source, temperature, pH, and inoculum density using the response surface methodology, obtaining an at least 33-fold increase in protease production [58]. Nonetheless, the principal difficulty in this phase is to find the most important factors to modify and their possible values for the optimization [59]. Therefore, testing is required afterward in order to empirically assess and confirm the optimization. Scale-up of the biomass and protein production Subsequently, it is necessary to scale up the whole process, including biomass generation and protein purification. Scale-up in bioreactors is an important process that allows obtaining a higher number of bio-products. In order to guarantee a good biotechnological process, it is recommended to utilize a volume ratio of 1:10 for a consecutive scale-up [60]. For example, to scale up a 100 L bioreactor, a 10 L culture is required. Biomass and product manufacturing at a larger scale must be as similar as possible to the laboratory scale. The aim of the scale-up is to replicate the behavior and results obtained in a lower scale, although the bioreactor design and mass transfer rate estimation is different in both scales. Also, variables such as agitation, aeration, and heat dissipation change with the size of the bioreactor, while temperature, pH, and viscosity remain constant. For example, the aeration rate and agitation must be reoptimized in the bioreactor because in a flask, oxygen is transferred by orbital agitation, while in a fermenter, it is transferred by air supply and the rotation speed of the impeller.

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An important criterion for scale-up is to maintain geometrical similarity, so that the behavior of the microorganism’s growth can be similar if two bioreactors maintain constant relations in their dimensions and lengths. This criterion should be applied whenever possible to design a larger-scale bioreactor [60, 61]. Another criterion to be considered for scale-up of aerobic fermentation processes corresponds to the volumetric oxygen transfer coefficient (kLa) [62]. The kLa coefficient allows measuring the gas-liquid transfer inside a bioreactor. This coefficient depends on different parameters such as aeration and agitation, chemical and rheological properties of the media, and the bioreactor geometry. This criterion can be determined through empirical or semiempirical formulas, which allows a correlated relationship among factors that affect oxygen transfer, like the impeller rotational speed, the volume of the fermenter, superficial air velocity, volumetric power input of un/ gassed system, and kLa [63]. There are many experimental methods for measuring kLa of bioreactors like the sulfite oxidation technique, gassing out, and dynamic methods [64, 65]. In order to use the kLa coefficient as a scale-up criterion, its value must be constant in different scales (e.g. 5 L, 50 L, and 500 L). An example of a scale-up process is shown in the work of Zhou and collaborators in 2018. They studied the growth of Streptomyces kanasenisi ZX01 and the production of a novel GP-1 glycoprotein. They successfully scaled up the fermentation from bench-scale to pilot scale until 500 L bioreactor, which was achieved using constant kLa coefficients [62]. Nevertheless, achieving reproducibility in each batch in relation with biomass and/or protein production still represents a challenge during the scaleup process. For this reason, it is necessary to have all parameters under control inside a bioreactor. If a batch is not reproducible, it must be eliminated, which translates into high economic losses. Enzyme purification scale-up There are three main purification methods depending on the technique or property of the enzyme to be purified. These are based on (a) the ionic properties of the enzyme, (b) the ability of the enzyme to get adsorbed, and (c) the differences in size of molecules [66]. Among the techniques based on the ionic properties of the enzymes, salting out is done by varying the pH of the solution or by the addition of chemical agents that carry out precipitation. The second technique used is ion exchange chromatography. Sophisticated systems for gradient formation and chromatographic resins (e.g. DEAE-sepharose, SP-sepharose) for separation are available, and they can be used in large columns in an automatized manner. Important factors such as column design, resin selection, and automation all play a role in simplifying the purification process, in some cases reducing the number of purification steps required to obtain a pure enzyme.

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Chromatography columns and systems are critical factors to the successful separation of the enzyme. The columns should be easy to pack and unpack, and they should provide a high linear velocity for maximum purification, which translates to maximum productivity [66, 67]. When selecting resins for large-scale purification, ionic exchange chromatography, hydrophobic interaction chromatography, and multimodal resins are common. In cases where ion exchangers cannot provide the required separation, hydrophobic interaction and mixed-mode adsorbents are generally used. Lately, periodic counter-current chromatography allows overloading of columns and continuous loading at the same time. Arranging two different modes of chromatography in series, operated with or without intermediate buffer conditioning, is another setup known as straight-through processing [68]. Techniques depending on the absorbing properties of the enzyme include adsorption and affinity chromatography [66, 67]. The first one is widely used for the recovery of extracellular enzymes. Affinity chromatography makes use of the interaction of a substrate with the enzyme. Among the techniques for purification depending on size of the enzymes are molecular sieve and gel permeation chromatography. Ultrafiltration and dialysis are used for enzyme purification as well as for concentration. Membrane based processes such as reverse osmosis and pervaporation are as well commonly used in purification procedures. For thermophilic enzymes, an easy approach to purify heat-resistant proteins is to denature the other proteins in the mixture by heating and then cooling the solution. Process validation is another key consideration in developing a large-scale purification strategy. This refers to establishing documented evidence, which assures that a specific process will consistently generate a product meeting its predetermined specifications.

4.4.4 Phase 4 – production phase Quality-controlled generation of biomass and enzyme production Shown below is a general example of recombinant enzyme production and quality control (Fig. 4.2). This process is divided in sequential activities with well-defined quality control points (QCPs) summarized in Tab. 4.1. Activation: The glycerol stock of the recombinant microorganism (e.g. E. coli BL21 harboring pET vector with the target gene inserted) must be activated. The activation is performed in sterile media (with antibiotic, if needed) from a cryogenic tube stored at −80°C. This phase finishes when the culture reaches an adequate optical density at 600 nm (i.e. 0.6–1). Then, the activated culture must be inoculated in a cotton-plugged flask with the appropriate medium, which is incubated in an orbital shaker set at the optimal temperature and agitation.

4.4 Direct exploration of enzymatic activities  

Seed culture preparation

Microorganisms activation

QCP1 Accepted?

YES

Biomass culture preparation

 113

YES QCP2 Accepted?

NO

NO Culture concentration

Protein solution preparation

NO

QCP5 Accepted?

Final product processing

YES

YES

QCP4 Accepted?

Protein purification

YES

QСР3 Accepted?

NO

NO

Product packaging labelling and delivery

Product documents

Fig. 4.2: Flowchart of the quality-controlled generation of biomass and enzyme production. Tab. 4.1: Quality control points during the production phase. QCP Number

Step

Control point

QCP1

Seed culture preparation.

Control of biomass growth from lab-scale bioreactor

QCP2

Biomass production

Control of biomass growth from a larger bioreactor

QCP3

Protein solutions preparation

QCP4

Protein Purification Final product processing

Control of enzyme activity and protein concentration of the solutions before loading onto the column. Control of specific activity of the eluted solution. Control of specific activity and protein concentration of the final product.

QCP5

Observations If the biomass yield is not as expected, the process should not be continued. If the activity is not accomplished with product specifications, most likely the final product will not accomplish the required specifications neither. Specific activity and concentration protein must be higher than formal specifications for the approval of the final product.

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Seed culture preparation: The culture obtained from the previous step is the inoculum required for the seed culture, which is then inoculated in a small bioreactor (e.g. 3 L) containing the appropriate sterile medium with antifoam and antibiotic if needed. Subsequently, the seed culture is grown under the optimal conditions of temperature, air supply, and agitation rate (cascade method can be used), with or without pH control, depending on the microorganism. At the end of this step, a fraction of the culture is centrifuged and the supernatant is eliminated. The pellet obtained is weighted in order to determine the wet biomass yield (QCP1). Biomass production: The seed culture obtained from the previous step is inoculated in an industrial bioreactor (i.e. 30 L) containing the appropriate sterile medium with antifoam and antibiotic if needed. This process can be performed using a peristaltic bomb with sterile silicone tubing. The optimal culture conditions of temperature, air supply, and agitation are specific for each recombinant microorganism and the optimal conditions should be the same that the optimal found for the small bioreactor. In this phase, it is important that the pH is controlled with buffer in the media or by using sodium hydroxide (NaOH) and hydrochloric acid (HCl) solutions. If the expression of the recombinant extremozyme requires the utilization of an inductor (e.g. IPTG), this must be added to the culture at the right time (i.e. normally at the middle exponential phase of the microorganism growth curve) and at its optimal concentration for the correct induction of the recombinant enzyme expression. Once biomass production has finished, it is again necessary to calculate the biomass yield of the process (QCP2). Culture concentration: The biomass obtained in an industrial bioreactor must be concentrated. This can be done using an appropriate filtration system that allows the reduction of the total volume for up to 90%. Then, the final volume must be centrifuged. In the case of expressing a recombinant protein extracellularly, the supernatant must be collected and further concentrated in order to proceed with the protein purification step. When working with intracellularly expressed recombinant enzymes, the cell pellet must be disrupted in order to obtain the crude extract necessary to continue with the protein purification step. If the cell pellet is not used, it should be stored immediately at −20°C. Protein solution preparation: In the case of proteins expressed extracellularly, further concentration is required in order to load an appropriate amount of protein onto the chromatography column. On the other hand, when working with proteins intracellularly expressed, the cell pellet obtained from the previous step must be resuspended in an appropriate buffer solution and the homogeneous mixture must be disrupted. This can be done using a cell disruptor equipment or by another mechanical process (e.g. French press, sonication). Because the solution-viscosity increases with the DNA released during the cell lysis, treatment with a DNAse enzyme is recommended.

4.5 Conclusions 

 115

The final protein solutions must be analyzed in terms of enzymatic activity (Units/ mL) and protein concentration (mg/mL) (QCP3). Protein purification: When the recombinant enzyme belongs to a hyperthermophilic or thermophilic microorganism, a heat-shock treatment (between 60 and 80°C) can be performed in order to eliminate some thermolabile proteins from the mesophilic host. The solution obtained is then ultra-centrifuged and the pellet is discarded. The protein solution is then loaded into a chromatography column for the correct purification of the target enzyme, using the specific conditions previously studied during the scale-up of the purification. Afterward, elution must be performed according to the type of resin used. Enzyme activity of the recombinant protein must be measured and comply with the expected values for its approval (QCP4). If the enzyme solution is accepted, it is filtrated in order to obtain an enzyme solution free of contaminants. Final product processing Often, but not always, enzyme products in powder format are preferred by certain markets due to their better stability and easier transportation compared with liquid ones. Powdered enzyme products are obtained through a lyophilizing process. The right package for an enzyme product must be studied according to the format (liquid or powder), stability, and storage conditions for each enzyme. The use of appropriate enzyme vials and their composition (e.g. glass, amber glass, plastic) must also be considered in order to guarantee correct enzyme storage. Then, one vial is randomly taken to measure the enzymatic activity of the final product. The specific activity and protein concentration must exceed the specification values for the final product approval (QCP5). It is a very common practice to exceed protein concentration and activity in the final product in case the enzyme suffers any issues during its distribution or storage. Thus, even though the enzyme may have lost activity, the final product is still in the accepted parameters reported in the Quality Control Certificate and Product Datasheet; if the product does not achieve its datasheet specifications, the whole batch must be discarded (Fig. 4.2). If the final product complies with the expected values, it is labeled and delivered with all the necessary paperwork (e.g. Analysis Certificate, Material Safety Data Sheet, and Certificate of Origin).

4.5 Conclusions The discovery of new enzymes and the development of new processes for enzyme production will drive future industrial innovation improving current industrial biocatalytic processes. In this new trend to replace chemical catalysis for biocatalysis,

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enzymes derived from microorganisms isolated from extreme environments have called the attention due to their novel properties allowing them to carry out reactions under nonstandard conditions, highly suitable for industry. However, as discussed in this chapter, the growth of extremophiles could be difficult and expensive due to their specific culture media requirements, low biomass yield, the need of specialized equipment, and high energy consumption, which translate to low quantities of native extremozymes. For these reasons, the development of recombinant microorganisms overexpressing extremozymes encoding genes, as well as their growth optimization for correct scaling-up process, is necessary for a more cost-effective enzyme production. In addition, it is worth to note that an adequate downstream processing and the establishment of QCPs at key stages of the bio-product production process must be implemented. Nevertheless, to definitely adopt extremozymes in industrial processes, the incorporation of modified procedures adapted for these biocatalysts have to be improved and developed combining functional, genomic, bioinformatic, and engineering approaches.

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Hans Jörg Kunte, Thomas Schwarz and Erwin A. Galinski

5 The compatible solute ectoine: protection mechanisms, strain development, and industrial production Abstract: Bacteria, Archaea, and Eukarya can adapt to saline environments by accumulating compatible solutes in order to maintain an osmotic equilibrium. Compatible solutes are of diverse chemical structure (sugars, polyols, amino acid derivatives) and are beneficial for bacterial cells not only as osmoregulatory solutes but also as protectants of proteins by mitigating detrimental effects of freezing, drying, and high temperatures. The aspartate derivative ectoine is a widespread compatible solute in Bacteria and possesses additional protective properties compared with other compatible solutes and stabilizes even whole cells against stresses such as ultraviolet radiation or cytotoxins. Here, it is our intention to go beyond a simple description of effects, but to depict the molecular interaction of ectoine with biomolecules, such as proteins, membranes, and DNA and explain the underlying principles. The stabilizing properties of ectoine attracted industry, which saw the potential to market ectoine as a novel active component in health care products and cosmetics. In joint efforts of industry and research, a large-scale fermentation procedure has been developed with the halophilic bacterium Halomonas elongata used as a producer strain. The development and application of ectoine-excreting mutants from H. elongata (“leaky” mutants) allow for the annual production of ectoine on a scale of tons. The details of the strain development and fermentation processes will be introduced.

5.1 Introduction Hypersaline environments with salt concentrations up to saturation of sodium ­chloride (approximately 300 g L−1) are mainly inhabited by microorganisms. Many hypersaline environments derived from seawater by evaporation and their ion composition are therefore similar to seawater. These so-called thalassohaline ­ ­environments comprise sodium and chloride as their main ions and have a neutral to slightly alkaline pH. Typical thalassohaline habitats are solar saltern crystallizer ponds, which can be found in tropical and subtropical areas around the world or the Great Salt Lake in Utah [1]. Athalassohaline hypersaline habitats exhibit a quite different pH and ion composition, and in the case of the Dead Sea, divalent cations such as Mg2+ and Ca2+ are dominating. Athalassohaline and thalassohaline environments provide harsh living conditions; however, both habitats harbor halophilic microorganisms, or halophiles, from all three domains of life: Archaea, Bacteria, and Eukarya. Halophilic microorganisms have developed two basically different https://doi.org/10.1515/9783110424331-005

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mechanisms and combinations thereof to cope with ionic strength and the considerable water stress, namely the “salt-in-cytoplasm” mechanism and the organic-osmolyte mechanism. Organisms following the salt-in-cytoplasm mechanism adapt the interior protein chemistry of the cell to high salt concentration [2]. The osmotic adjustment of the cell can be achieved by raising the salt concentration (KCl) in the cytoplasm according to the environmental osmolarity [3]. In contrast, microorganisms applying the organic-osmolyte strategy keep their cytoplasm, to a large extent, free of KCl and the design of the cell’s interior remains basically unchanged. Instead, organisms of this group accumulate uncharged, highly water-soluble, organic compounds in order to maintain an osmotic equilibrium with the surrounding medium. The organic-­ osmolyte mechanism is widespread among Bacteria and Eukarya and also present in some Archaea [4–6]. Organic osmolytes are of diverse chemical structure comprising different types of sugars (e.g. trehalose), polyols, amino acids (proline), and their derivatives (ectoine, glycine betaine) and are accumulated inside the cell either by de novo synthesis or by uptake from the surrounding environment. These nonionic, highly water-soluble molecules do not disturb the metabolism, even at high cytoplasmic concentrations, and are thus aptly named compatible solutes [7]. Compatible solutes are beneficial for bacterial cells not only as osmoregulatory solutes but also as protectants of proteins by mitigating detrimental effects of freezing, drying, and high temperatures [8]. Ectoine (1,4,5,6-tetrahydro-2-methyl-4-pyrimidine carboxylic acid) [9], which is a widespread compatible solute in Bacteria [10–12], but also – albeit rare – in Archaea and Eucarya, possesses additional protective properties compared with other compatible solutes such as glycine betaine and proline and stabilizes even whole cells against stresses such as ultraviolet (UV) radiation, inflammation caused by nanoparticles or cytotoxins [13–18]. The properties of ectoine attracted industry, which saw the potential to put a novel protective compound for healthcare products on the market. Ectoine can be synthesized chemically, but not for an acceptable price due to the high costs of the precursors. This led to the development of a more competitive biotechnological production of ectoine with halophilic bacteria. Halophilic microorganisms have been used for food fermentation in the Far East since centuries and are of importance for food industry in the production of flavoring agents [19]. However, besides food industry, there are only a few biotechnical applications with halophiles that are successfully in use with industry. Most biotechnological applications exploiting halophiles and their products are still under development, such as the application of bacteriorhodopsin as a holographic storage material, the production of poly-β-hydroxyalkanoate and extracellular polysaccharides by Haloferax mediterranei, and novel enzymes (e.g. restriction enzymes, proteases) from different halophilic Archaea and Bacteria. The only commercially successful applications with halophiles today are the production of β-carotene and ectoines. The β-carotene is produced by different species of the unicellular green alga Dunaliella and the first biotechnical production plant was already operational in the late 1960s in the former USSR [20]. Today, β-carotene is produced around the world [21] and is used

5.2 Molecular interaction of ectoine with water and biomolecules 

 123

as a food-coloring agent, as pro-vitamin A (retinol), and as an additive in cosmetics. The second biological production process with halophiles that is successful in the market is the above-mentioned production of the cell protectant ectoine. Ectoine, the flagship of “extremolytes” [22], is synthesized on a scale of tons by the company bitop (Dortmund, Germany) in a process with the halophilic γ-proteobacterium Halomonas elongata used as producer strain. Its protective properties as stabilizer of enzymes, membranes, DNA, and whole cells make ectoine a valuable compound, which is marketed in healthcare products and cosmetics. Our purpose here is to review the different aspects of the biotechnology of ectoine. In particular, the present chapter will illustrate the mechanism of ectoine in stabilizing biological structures and explain the development of natural producer strains and their use in industrial fermentation processes.

5.2 Molecular interaction of ectoine with water and biomolecules and its effect on solution properties Ectoines, like other compatible solutes, are selected by nature as a protection against environmental stresses. As such, they also improve the stability of biomolecules. The protein- and enzyme-stabilizing phenomena of ectoine were among the first observations and have been reported by many authors and been included in many reviews on cell stress protection [23, 24]. Here, it is our intention to go beyond a simple description of effects, but to depict the molecular interaction of ectoine with biomolecules, such as proteins, membranes, and DNA, and explain the underlying principles that make it an excellent excipient for a variety of applications. In order to achieve this, ectoine must be seen in context with other natural stabilizing compounds, the most prominent of which are N-methylated derivatives such as trimethylamine N-oxide (TMAO), glycine betaine, proline, sugars, and polyols. Although one needs to keep in mind that stabilization phenomena reported by different authors might, to a certain degree, be specific for the system under investigation, we will try and extract general properties that are (more or less) characteristic for all compatible solutes and pinpoint special phenomena that distinguish ectoine from other common solutes, in particular glycine betaine.

5.2.1 Proteins Preferential exclusion The pioneering work of the Timasheff group of the 1980s has shown experimentally that the compatible solutes (protecting osmolytes) under investigation were preferentially excluded from a protein’s hydration shell. Due to their preferential exclusion from the protein surface, the phenomenon of protein stabilization could therefore be

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explained as the result of the tendency to minimize the surface area [25–27]. Although ectoines were not included at the time of the study, later work, using different techniques, has confirmed the exclusion phenomenon for ectoine [28]. However, which forces are responsible for the observed exclusion and subsequent stabilization is only slowly becoming clear. Osmophobic effect The work of the Bolen group has identified the peptide backbone as the major player for binding/repulsion effects, and the subsequent protein stabilization [29, 30], and named this phenomenon the osmophobic effect [31]. Using diketopiperazine, a cyclic dimer of glycine, as model compound for the peptide backbone, they were able to demonstrate that all compatible solutes tested were characterized by a positive transfer free energy (ΔGtr) of backbone into osmolyte solution, which means that the solute-containing solution is an unfavorable solvent for the backbone. Subsequently, an apparent correlation was observed between the measured ΔGtr and fractional polar surface of the compatible solute. This allowed establishing a model in which the interaction energy of the solute with the peptide backbone depended on the polarity of the solute and the fractional polar surface area [32]. Despite the simplicity of this approach, neglecting other types of interaction, the model correlated very well with experimental values of ΔGtr and emphasized the validity of the underlying concept. As osmolyte backbone interactions become increasingly favorable with osmolytes becoming more polar, the stabilizing properties of an osmolyte are predicted to improve with increasing hydrophobic surface area while at the same time still maintaining solubility in water. The quaternary ammonium group (as in TMAO, glycine betaine etc.), but also the 2-methyl pyrimidine group of ectoines, seems to be especially important as they convey a hydrophobic increment while at the same time avoiding hydrophobic fusion due to the repulsion caused by the positive charge. Therefore, it is very likely that protein stabilization by ectoines is based on similar molecular mechanisms as those described for other nitrogen-containing zwitterionic compatible solutes, such as TMAO, glycine betaine, and proline [33]. This model for osmolyte-induced folding/unfolding of proteins [34] was later partly revised when the influence of individual side chains on the accessible surfaces of the backbone and activity coefficients were accounted for [35]. This proved that the contribution of the side chains had been underestimated before and that, in case of urea, backbone and side chains contribute approximately the same to its protein denaturation effect. From this, one must conclude that peptide side chains are not completely irrelevant, albeit much less important than the backbone [30, 36]. In the case of ectoine, a possible interaction with aromatic amino acid side chains is predicted, due to the structural arrangement (tryptophane box) of the binding pockets of ectoine binding-proteins [37–39]. On the basis of these observations, one can conclude that

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a solvent containing compatible solutes becomes a poorer solvent for the peptide backbone and for most of the side chains. By virtue of reducing its interaction with the solvent, intrachain hydrogen bondings are strengthened and secondary structures stabilized [40]. Conformational consequences As the α-helical core in particular resembles a conformation in which exposure of the backbone to the solvent is minimal, one would expect compatible solutes to strengthen this formation. This concept has gained support by the finding of Bourot et al. [41] that a metabolically inactive diaminopimelate decarboxylase (Ser 384 Phe mutation) is reactivated in the presence of compatible solutes like glycine betaine in vitro and in vivo. As demonstrated by the authors, the molecular cause for inactivity can be traced back to a loss of α-helical structure, which is apparently recovered in the presence of compatible solutes. The authors conclude that compatible solutes may assist protein folding in a “chaperone-like” manner. Similarly, in a study on alanine-based model peptides, trimethylamine-N-oxide (TMAO) excelled as an osmolyte, which induced helix formation [42] as predicted by the osmophobic effect hypothesis. A Raman study of the amide I region of the DNA-binding gene-5-protein (G5P) also revealed slight conformational changes in the presence of 1 M ectoine, i.e. an increase in α-helix at the expense of β-sheet and β-turn [43]. It is our opinion that compatible solutes may be best described as low-molecular mass solutes enhancing the stability of secondary structures (in particular α-helical elements) of the peptide backbone (“backbone chaperones”). For ectoine, this view has found experimental support in atomic force microscopy (AFM) studies, which showed a general tendency (like with other compatible solutes) to enhance intramolecular forces and “stiffen” the structure of a protein, at least in case of fibronectin and the membrane protein bacteriorhodopsin [44, 45]. Preliminary results on protein aggregation studies seem to indicate that compatible solutes (including ectoine) may also prevent pathogenic protein aggregation [16] and, in particular, β-sheet-based amyloid formation as observed in neurodegenerative disease [46].

5.2.2 Membranes Concerning the interaction with other biomolecules, as for example lipids of the cytoplasmic membrane, research on native fluid systems (as opposed to dry stabilization of lipsomes) is still at the beginning. Experimental evidence of the Galla group using the Langmuir film-balance technique, epifluorescence microscopy, and atomic force microscopy (AFM) revealed striking effects of ectoines on the fluid-rigid domain

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distribution. The presence of ectoines evoked smaller and more numerous domains, thus expanding and fluidizing lipid monolayers [47]. Therefore, ectoines appear to have an effect on the line tension by reducing the interfacial energy at the edge of the domains and on the mobility of the head groups. This effect was shown to be opposite to that of urea [48], which suggests that the counteractive function toward denaturants also applies to the fluid membrane system. Membrane interaction studies with ectoines were subsequently extended to the domain structure of artificial lung surfactant films, where their positive effect on biophysical compression/expansion behavior was confirmed [49]. In addition, infrared reflection-absorption spectroscopy indicated that an interaction with the phospholipid head groups (at higher concentration) cannot be excluded, at least not for hydroxyectoine [49]. However, it is very likely that the indirect effects of modulated solvent properties (see Chapter 5.2.5) may also play a part. It remains, for example, an interesting option that changes in the dielectric constant of the solvent will have a positive effect on the hydration and subsequent interspacing of the polar (zwitterionic) phospholipid head groups. Work on the molecular interaction of ectoines on lipid films has subsequently been expanded to tear fluid bilayers, which consist of a polar sublayer (in contact with the tear fluid on the corneal epithelium) and a nonpolar upper layer in contact with the air. This structure seals the tear fluid from the air and reduces evaporation. In “dry eye syndrome,” which is caused by a dysfunction of the meibomian gland, the rigidified lipid layer raptures easily, leading to increased rate of evaporation of the tear fluid. It has been shown for both the natural meibomian film with phosphatidyl choline as major compound and artificial lipid films that ectoine fluidizes the polar monolayer in contact with the tear fluid by increasing the area occupied by the polar lipid head groups, thus decreasing the rigidity of the film [50, 51]. This fluidizing effect of ectoine has since found an application in eye drops. A subsequent study on giant unilammelar bilayer vesicles of phosphatidyl choline lipids proved an increase in the disordered (ld) phase at the expense of the liquid ordered (lo) phase in the presence of ectoine (and hydroxyectoine), implying a relative increase in fluidity [52]. As N-methylated compatible solutes (TMAO, glycine betaine, etc.) have so far not been included in such fluid membrane studies, it is not possible to conclude whether the observed effect on membranes is typical for ectoines or a general feature of compatible solutes. Neutron scattering studies on the purple membrane of Halobacterium salinarum have also indicated an exclusion of ectoine from the hydration layer of the membrane surface [53]. However, for this system, the authors came to the (surprising) conclusion that the molecular dynamics of the membrane are not affected by the presence of ectoine. In this context, one must bear in mind the special composition of the purple membrane [54], which consists of mostly anionic phospholipids and a large proportion of protein, and the fact that the experimental conditions were far from physiological (progressive drying of 1 M ectoine with purple membrane).

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5.2.3 Nucleic acids A review on the effect of compatible solutes on nucleic acids [55] summarizes all potential interactions of ectoines and other compatible solutes, including indirect countercation effects through the modulation of electrostatic properties as described by Flock et al. [56] and Houssier et al. [57]. Already in 1993, Rees et al. [58] reported that glycine betaine decreased the melting temperature of DNA. In addition, the authors observed (using DNA of different GC content) that the effect was more pronounced with DNA of high GC content and caused an elimination of the (natural) differences in melting temperature. This observation has led to the application of glycine betaine (and other cosolutes) as polymerase chain reaction enhancers for GC-rich templates [59]. It was also shown that the effect of glycine betaine was greatly enhanced when the methyl groups were synthetically replaced by more hydrophobic moieties (i.e. butyl residues) [60]. A similar effect (improved efficacy) has also been demonstrated for the more hydrophobic ectoine derivative homoectoine [61, 62]. As regards the potential interaction of glycine betaine with DNA, Hong et al. (2004), using vapor pressure osmometry, reached the conclusion that this solute is excluded from the anionic DNA surface and that the hydration of the phosphate oxygen surface involves at least two layers of water [63]. They postulated that glycine betaine is a specific GC denaturant because of its favorable interaction with G and/or C in the single-stranded state (ssDNA). This view was corroborated by work on osmolyte effects on RNA secondary structure stating that glycine betaine (among other osmolytes like TMAO and proline) preferentially accumulated at the surface of ssDNA, when bases are exposed to the solvent, especially G and C [64]. At the level of tertiary RNA structures, however, there seemed to be a similar exclusion phenomenon from the backbone phosphate as with the peptide backbone of proteins (at least for TMAO) [65]. The interaction of betaine with DNA (in comparison to urea) was further investigated using a range of different techniques (FT-IR spectroscopy, differential scanning and isothermal titration calorimetry, and molecular dynamics simulations) [66] on calf thymus DNA in a buffer-free system. On the basis of their IR data, the authors come to the conclusion that betaine has no direct specific interaction with calf thymus double-stranded DNA (dsDNA) but with increasing concentration (0.5–3.5 M) shifts the conformational equilibrium from B- to A-DNA. Molecular dynamic simulations seem to support this view of relative exclusion from the surface (albeit with a small aggregation in the second solvation sphere, i.e. at distances greater than 0.4 nm) and suggest that DNA undergoes a compression along the helix axis, as would be expected for the B-to-A conformational change. Nevertheless, there also appears to be a small trend toward specific interactions with the bases of dsDNA, in particular via G-C or C-G in the DNA sequence. Upon denaturation, the biggest increase in the number of hydrogen bonds was observed for betaine with guanine residues, which corroborates the experimental findings of a GC-dependent melting point reduction.

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As ectoine, contrary to TMAO and betaine, is a good hydrogen acceptor as well as a donor, one would expect a stronger interaction with nucleic acid bases. A spectroscopic study using synthetic double-stranded DNA (20 base pairs, 65% GC) in phosphate buffer (plus NaCl) and increasing concentrations of ectoine (up to 3 M) revealed a linear concentration-dependent decrease in melting temperature [67]. The same authors also conducted molecular dynamics simulations on short single-stranded DNA hairpin (seven nucleotides) and a 24-base pair duplex B-DNA structure in a buffer-free system with various concentrations of the cosolute. Against expectations, evaluation of the radial distribution functions displayed a concentration-dependent effect. At low concentrations (up to 1 M), ectoine was attracted strongly to all DNA conformations, albeit more though to single-stranded DNA. At higher concentrations, however, ectoine increased in the bulk solution, as expected for a preferential exclusion solute [67]. The authors proposed that ectoine partially replaces water molecules around the DNA as a consequence of the observed preferential binding up to the point when hydration shells are saturated. Ectoine’s unfolding impact on DNA was interpreted as a consequence of its higher affinity to ssDNA. The puzzling observation of preferential binding must be seen in the context of this artificial model system where the negative charges of the phosphate backbone are “neutralized” with sodium counter ions, but the possible impact of other buffer salts is neglected. It therefore remains to be shown whether ectoine is still attracted to the backbone of dsDNA under physiological conditions when counterions are abundant. In a study investigating the UV-protective effect of ectoine on plasmid DNA, it was observed that, contrary to expectations, single-strand breaks were enhanced at 500 mM ectoine in pure water. In addition, the effect of UV-A irradiation was enhanced in combination with ectoine [68]. As the stability of plasmid DNA was, however, unaltered in a buffered solution (pH 7.5) containing ectoine, it was hypothesized that the ectoine effect in pure water may be caused by locally increased acidity and as such unphysiological. The authors speculated that the damage-enhancing effect during UV-A irradiation (365 nm) may be caused by a photosensitizing action of high ectoine concentrations in close vicinity of DNA [68]. When plasmid DNA in water was directly irradiated in a special sample holder with electrons of a primary kinetic energy of 30 keV, indirect effects of secondary particles (OH-radicals, ions, and low-energy electrons) were identified as major cause of single-strand breaks [69, 70]. In a subsequent publication, the authors analyzed the protective effect of ectoine in PBS buffer, which had its maximum at 0.8 M (no damage). In order to explain this effect, they argued that ectoine may protect by passively displacing water surrounding the DNA, and in doing so may reduce the production of secondary damaging agents in its vicinity [71]. Alternatively, the vibrational behavior of strongly structured water in the vicinity of this kosmotrope could be (at least partly) responsible by increasing the probability of inelastic scattering (and energy loss) of low-energy electrons. Eventually, OH radical scavenging of ectoine was also identified as a possible contribution to the protection of DNA against radiation [71, 72].

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A recent paper by Hahn et al. [73] showed that during irradiation with UV-C (266 nm), ectoines protected against single-strand breaks of plasmid DNA in pure water, whereas in buffer (i.e. under more physiological conditions), ectoines protected against base damage (cyclobutane pyrimidine dimer formation). In an attempt to identify the mode of interaction with nucleic acid, the same authors conducted small-angle X-ray scattering, Raman spectroscopy, and surface plasmon resonance (SPR) studies, the latter with surface tethered ssDNA (dT25 oligomers). SPR clearly proved concentration-dependent binding of ectoine to ssDNA with a maximum of 1.44 ectoine molecules per nucleotide at the highest measured concentration (1 M). For buffer-free dsDNA (Herring sperm), however, Raman spectroscopy indicated changes in the hydrogen bondings with both ectoine and DNA, while X-ray scattering suggested changes in the conformation of DNA, in both cases for low ectoine concentrations (40 g dry weight/L corresponding to >10 g/L ectoine). The bacterial milking process exploits the ability of H. elongata to release ectoine in response to dilution stress to the medium. Cells of H. elongata are grown in a fed-batch fermentation process at a salinity of 10% NaCl (1.7 M) until a high cell density has been reached. Then, an osmotic down shock from 10% to 2% NaCl is applied. As a result, approximately 80% of the cytoplasmic ectoine is released to the culture medium. The excreted ectoine is then recovered from the medium by cross-flow filtration and further purified by cation exchange chromatography and crystallization. After harvesting, the filtered cell mass is reused and brought back into high saline (10% NaCl) culture medium for another round of ectoine synthesis. Within 10 hours, the bacterial milking can be repeated. A significant increase in productivity could be achieved by replacing the fed-batch process with a continuous fermentation [130]. The bacterial milking procedure was the first process implemented in industry for the large-scale production of ectoine but has been replaced by a more efficient process relying on strains of H. elongata which excrete ectoine.

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5.3.4 The “leaky” mutant procedure (exploiting a mutant which excretes ectoine into the medium) Microorganisms do not rely entirely on de novo synthesis of compatible solutes. They are able to take up compatible solutes or precursors from the surrounding environment, if available. To allow for this uptake, microorganisms must be equipped with a set of specific transporters functioning at high osmolarity and high ionic strength, conditions that often inhibit transporters for nutrient uptake. These transporters facilitate a far more economical accumulation of compatible solutes compared to de novo synthesis [122]. H. elongata is equipped with a set of different compatible solute transporter [123], but only one transporter facilitates the (osmoregulated) uptake of ectoine namely the tripartite ATP-independent periplasmic transporter (TRAP-T) [131, 132] TeaABC [133]. TRAP-T are widespread in organisms from the bacteria and archaea domain. Unlike the ABC transporters that are fuelled by ATP hydrolysis, TRAP-T are secondary transporters. Reconstitution experiments with the TRAP-T SiaPQM revealed that Na+ is the coupling ion for transport. This finding is supported by the fact that TRAP-T can be found predominantly in marine bacteria, which prefer to utilize Na+ as a counter-ion due to its abundance. The actual transporter TeaABC consists of three proteins, a large transmembrane protein (TeaC, Helo_4276), a small transmembrane protein (TeaB, Helo_4275), and a periplasmic substrate-binding protein (SBP; TeaA, Helo_4274) with a high affinity for ectoine [38, 134]. In addition, a regulatory ATP-binding protein TeaD (Helo_4277) is influencing the activity of TeaABC. The larger transmembrane protein is thought to catalyze the actual transport reaction and is responsible for energy coupling, most likely through the sodium motive force. The small membrane protein is not involved directly in the transport reaction and its exact function is still unknown. Since the small subunit is almost always encoded upstream of the gene for the large subunit, a chaperone-like function in the folding of the large membrane protein is suggested. Another hypothesis brought forward is that the small subunit might be needed to interact with the SBP for transport. TeaD is an ATP-binding protein and functions as a negative regulator of the TeaABC transporter [135]. However, the exact role of TeaD in regulating TeaABC is still unknown. Because osmoregulated transporters such as TeaABCD are exposed to both the high-salt environment and the cytoplasm, it was hypothesized that these systems would also function as sensors for osmotic changes. This was shown to be the case for different compatible solute transporters. In addition, compatible solute transporters must be integrated (directly or indirectly) in the regulation of the cell’s compatible solute synthesis, because the uptake of external solutes results in an immediate decrease in the concentration of compatible solutes synthesized by the cell [123]. In some bacteria, transporters for compatible solutes are not restricted to the accumulation of external osmoprotectants but are also functioning as a salvage system for compatible solutes leaking out of the cell [136]. That was shown to be true for

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H. elongata [133, 137]. Mutation of the ectoine transporter TeaABC (∆teaC) resulted in a mutant excreting ectoine to the medium. Quantifying the excreted ectoine and the cytoplasmic ectoine revealed that the TeaABCD transporter mutant is both an ectoine leaky mutant and an overproducer for ectoine [138]. Why ectoine is leaking out the cell when the specific transporter for ectoine is mutated is not quite understood yet. Similar observations have been recorded for the cyanobacterium Synechocystis sp. strain PCC 6803, which synthesizes glucosylglycerol as its main compatible solute [136, 139], and Bacillus subtilis, which synthesizes proline as compatible solute [140]. The mutation of ggtA, a gene encoding a subunit of an ABC transport system mediating the uptake of glucosylglycerol in Synechocystis sp., created a mutant leaky for glucosylglycerol, while the mutation of the osmoregulated proline transporter opuE in B. subtilis created a leaky mutant for proline. However, the mutation of the transporter in B. subtilis did not create an overproducing strain as observed for H. elongata, but quite contrary, a mutant that synthesizes less proline than the parental strain. There is evidence that the loss and the recovery of ectoine by the TeaABC transporter might be an elegant mechanism to regulate the cytoplasmic ectoine content in H. elongata [141]. As mentioned above, compatible solute transporters have to be linked to ectoine synthesis in order to shut down ectoine synthesis when external solutes are taken up. The same must be true for TeaABCD in order to regulate ectoine synthesis when external ectoine is transported into the cytoplasm. This implies that any ectoine, regardless of its origin (even ectoine lost from the cell), will have this negative regulatory effect on the synthesis of ectoine. It is therefore possible that ectoine transport could serve as a signal for the regulation of ectoine synthesis. Increasing the ectoine concentration by de novo synthesis will lead to water influx and an increase in turgor pressure of salt-stressed cells. Assuming that the efflux of ectoine via an export channel is triggered by a signal such as membrane stretch, ectoine will be released to the periplasm if the turgor pressure reaches a certain threshold. Transport of exported ectoine back into the cytoplasm by the activated TeaABCD system will down-regulate ectoine synthesis. The proposed regulation mechanism would allow for an oscillation of the cytoplasmic ectoine level closely above and below the threshold needed to open the solute-specific export channels. As a consequence, in cells with a functional TeaABC transporter, ectoine will be cycled between the outside (periplasm) and the cytoplasm. The described model is supported by the findings made with other bacteria (for more details, see below, “Mechanisms of ectoine export from H.  elongata cells”). It is important to point out the differences in the transport mutants of B. subtilis and H. elongata. The intracellular proline content in the opuE mutant from B. subtilis is considerably lower than that of its parental strain, and as a consequence, the mutant is salt sensitive. In contrast, teaABC mutants of H.  elongata have the same intracellular ectoine content and display the same salt tolerance as the wild-type cells [133]. As explained before, teaABC mutants of H.  elongata are overproducing

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mutants, which indicates a regulatory connection between the release and uptake of ectoine. These differences suggest a different osmoregulatory mechanism in B. subtilis compared to H. elongata in controlling the compatible solute content. Employing the ectoine overproducing “peeing” mutant of H. elongata for largescale industrial production of ectoine offers a number of advantages over the previously used bacterial milking procedure. Here, the production of ectoine is completely uncoupled from intracellular accumulation of ectoine; therefore, the yield in g/g on carbon source is substantially increased. This means that also the product titer is no longer limited by the biomass density that can be achieved. The process can be performed in continuous culture, but also in the simpler fed-batch or batch mode with the advantages of increased robustness and straightforward scalability. Also, the amount of salt in the process can be reduced to the level needed for the activation of ectoine biosynthesis enzymes, decreasing the salt burden that needs to be separated in the downstream processing. Furthermore, the number of downstream processing operations can be reduced, since no more osmotic down shock and second filtration step is needed. Altogether, the “leaky” production strain allows for a process more similar to the highly efficient fermentation processes employed for the manufacture of amino acids like lysine.

5.3.5 Mechanisms of ectoine export from H. elongata cells When bacterial cells are exposed to hypoosmotic shock (e.g. rainfall), MS channels allow for the unspecific and sudden efflux of soluble solutes from the cytoplasm in order to avoid immediate cell disruption. H. elongata possesses four MS channels of the MscS type, of which all are crucial for survival when facing hypoosmotic stress. They are also the export route for ectoine upon hypoosmotic shock and the underlying mechanism for the bacterial milking process. However, how ectoine is exported under steady-state conditions from osmotically adapted cells is still unknown. There are, in principle, four different ways by which compatible solutes can exit the bacterial cell, namely by (i) passive diffusion across the membrane, (ii) reversal of compatible solute uptake systems, (iii) efflux via (more or less unspecific) channels including MS channels (channel model), and (iv) efflux via carrier systems with either broad or narrow substrate specificity (carrier model). Passive diffusion across the membrane is no plausible mechanism in our view for the release of bacterial compatible solutes. There is also, to our knowledge, no support by proven examples for the release of compatible solutes by reversal of uptake transporters, and this can be ruled out for H. elongata as the only transporter for ectoine uptake, TeaABC, is absent in the leaky mutant [133]. Börngen et al., working on Corynebacterium glutamicum, presented results that argued for a mechanism of fine-tuning the internal compatible solute concentration of glycine betaine by cooperation of an active uptake via transporter BetP and

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a passive, but regulated, efflux via MS channels [142]. For releasing the compatible solute glycine betaine, C. glutamicum employs MscCG, a MS channel of the MscS-type. MscCG is the major export route for glutamate in C. glutamicum, and in addition to its osmotic function, it serves as a metabolic valve [143]. Compared to canonical MscS, MscCG is structurally different, which results in a different gating behavior with a lower activation threshold and slow closing [144, 145]. Whether the export via MS channels in bacteria is a widespread mechanism in fine-tuning the cell’s compatible solute concentration is still debated. A channelindependent export of compatible solutes was described for B. subtilis and E. coli. In B. subtilis, none of the known MS channels of the MscL and MscS type participated in the export of the endogenous compatible solute proline [146], and in osmotically adapted E. coli cells, hydroxyectoine export is independent of MS channels as well [147]. The recent findings by Vandrich et al. [129] draw a similar picture for H. elongata and revealed a MS channel-independent export of ectoine from osmotically adapted cells. They compared ectoine excretion of a “leaky” mutant, which is devoid of all four MS channels, with the parental “leaky” strain, which is equipped with all four operational MS channels. Both strains were grown at low (0.7 M NaCl) and high (2 M NaCl) salt growth media. During a 10-hour growth, the MS channel mutant exported indeed less ectoine, but the excretion of ectoine was only 19% (at 2 M NaCl) and 21% (at 0.7 M NaCl) lower than for the parental strain. Even though MS channels of H. elongata are an export route for ectoine, they are only of minor importance for this process. Approximately 80% of the ectoine was released from the cell via a major, yet enigmatic, pathway. Specific efflux systems for various types of compounds, in particular amino acids, are already known for different microorganisms [148–150]. Hence, H.  elongata might release ectoine via a carrier system, which is either a general compatible solutes exporter or a more specific system for the export of ectoine and, eventually, hydroxyectoine. Still the question remains on how an efflux system for compatible solutes is linked to osmolarity and can sense the internal solute concentration. For Dickeya dadantii (formerly known as Erwinia chrysanthemi), a putative sensor protein was described that enables the cell to specifically measure the internal glycine betaine concentration and the extracellular salt concentration [151]. Inactivation of this sensor, which shares some similarities with MscK, resulted in loss of glycine betaine from the cell but not of other compatible solutes.

5.3.6 Blocking ectoine degradation – the “super-leaky” mutant Ectoine is accumulated as an osmolyte in H. elongata. However, ectoine can also be used as both a carbon and nitrogen source, and when offered as a nutrient, ectoine

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still serves as compatible solute. In a combined effort of research and industry, the genome of H.  elongata was sequenced [12, 152]. Genome analysis and mutagenesis experiments led to the discovery of the pathway for ectoine degradation [12]. The degradation of ectoine (doe) to aspartate comprises four steps and proceeds via hydrolysis of ectoine to Nα-acetyl-L-2,4-diaminobutyric acid, followed by deacetylation to diaminobutyric acid. In H. elongata, diaminobutyric acid can either flow off to aspartate or reenter the ectoine synthesis pathway, forming a cycle of ectoine synthesis and degradation. The ectoine hydrolase DoeA, which catalyzes the first step in ectoine degradation, is a 399-aa protein (44.9 kDa, pI 5.0) that belongs to the peptidase-M24 family. Within that family, DoeA is similar to creatinase (creatine amidino-hydrolase), which catalyzes the hydrolysis of creatine to sarcosine and urea. The essential product of ectoine hydrolysis catalyzed by DoeA is Nα-acetyl-L-2,4-diaminobutyric acid (Nα-Ac-DABA), which is the substrate for the subsequent catabolic enzyme DoeB. In recombinant E. coli expressing doeA, also Nγ-Ac-DABA was detectable. Whether DoeA catalyzes the formation of both isoforms (Nα-Ac-DABA and Nγ-Ac-DABA) or if the cleavage produces only one isoform that is subsequently converted into the corresponding isomer by an acetyltransferase is not quite clear yet. The Nα-acetyl-L-2,4-diaminobutyric acid deacetylase DoeB is 342-aa protein (36.6 kDa, pI 4.6) and closely related to proteins of the succinyl-glutamate desuccinylase/aspartoacylase subfamily, which are part of the M14 family of metallocarboxypeptidases. DoeB deacetylates Nα-Ac-DABA to L-2,4-diaminobutyric acid (DABA), which can serve as substrate for the catabolic transaminase reaction DoeD. DoeD is a 469-aa enzyme (50.8 kDa, pI 5.6) that belongs to the PLP-dependent aspartate aminotransferase superfamily. DoeD catalyzes the formation of aspartate-β-semialdehyde, which finally can be converted to aspartate by dehydrogenase DoeC (493 aa, 53.1 kDa, pI 4.8). The genes encoding the ectoine degradation pathway are located together on the chromosome of H. elongata. The genes doeA and doeB are organized as an operon together with a third gene called doeX. The doeCD components are located adjacent to doeABX but are not part of the doeABX operon. Promoter-mapping analysis revealed the presence of a σ70-dependent promoter in front of the doeABX operon. The doeX gene encodes for DNA-binding protein belonging to the Asn/Lrp family of DNAbinding proteins and binds to a 46-nt sequence located directly upstream to the doeA start codon. The regulatory impact of DoeX on the doeAB expression is still unknown. As indicated above, DABA can be degraded to aspartate or reenter the ectoine synthesis pathway via EctA and EctC. In this way, degradation and synthesis of ectoine are directly connected, forming a cycle that is powered by the acetylation reaction catalyzed by EctA. However, this cycle appears to be futile, in which ectoine can be synthesized and degraded simultaneously, resulting in a net conversion of acetyl-CoA into acetate. Acetate can be reconverted to acetyl CoA by acetate:CoA ligase (AMP-forming; EC 6.2.1.1). Therefore, if an ectoine synthesis and degradation cycle were active, it would result in an increased cost of two ATPs per turn for ectoine

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synthesis. Schwibbert et al. [12] showed that such a cycle is an elegant mechanism to control the speed of change in internal ectoine concentration as a response to external changes in osmolarity. It is known that the turnover time of a metabolite, defined as the ratio between its concentration and the flux through it in the steady state, is a good indicator of the timescale of its transient responses. Metabolites with a high turnover tend to complete transitions faster than those with a low turnover. Thus, by keeping a flux through the synthesis/degradation cycle, the cell can achieve fast changes in ectoine levels to quickly respond to changes in external osmolality. Based on this model, a disruption of the ectoine degradation pathway should lead to a lower ATP load for cells synthesizing ectoine and at the same time in higher ectoine productivity. In fact, deletion of the doeA gene in an ectoine excretion mutant (KB2.11; ∆teaABC) resulted a strain of H. elongata with a three times higher productivity compared to the leaky mutant (Kunte, unpublished data). This new strain with a blocked ectoine degradation pathway and thereby increased ectoine excretion offers new options for the large-scale production of ectoine.

5.4 Products The first commercial use of ectoine was as a skin care ingredient [153, 154], and this application still plays a major role, especially in sun protection and antiaging products, where ectoine is widely used [23, 155, 156]. Recently, the use of ectoine in healthcare products has become of increasing importance. Starting with the work of Buenger and Driller [157] and Grether-Beck et al. [158], which showed that ectoine is able to inhibit the early UV-A radiation-induced ceramide signaling response in human keratinocytes, the mitigating effect of ectoine on inflammatory conditions of human skin was examined further. Pretreatment of keratinocytes with ectoine reduced the number of sunburn cells, prevented the decline in Langerhans cells [159], and decreased UV-induced DNA single-strand breaks [160]. In vitro studies on ectoine’s beneficial effects for human skin are confirmed by clinical trials on skin aging [161]. Here, it could be demonstrated that a formulation with already good skin care properties could be significantly improved by the addition of 2% ectoine, which led to superior skin hydration, skin elasticity, and skin surface structure. The mitigating effect of ectoine was not only observed in skin but also in inflammatory conditions of other epithelia. For lung epithelia, it was shown that ectoine protects against nanoparticle-induced airway inflammation [14, 162]. For nasal and eye epithelia in allergic conjunctivitis, ectoine-containing nasal spray and eye drop products relieved effectively the hallmark symptoms of rhinoconjunctivitis, with treatment effects similar to those of antihistamines, steroids, and leukotriene modifiers, but with virtually no side effects [163]. Based on this, a wide range of ectoine-based medical device products for the treatment of allergies (allergy nasal sprays, nasal douches, mouth and throat sprays,

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 5 The compatible solute ectoine

lozenges, inhalation solutions, wound healing products, dermatitis creams, ear sprays and eye drops), skin inflammatory conditions like atopic dermatitis, dry eye and dry nose, and rhinosinusitis have been developed, have been successfully tested in clinical trials [164], and have entered the market. Due to the excellent safety profile in combination with clinically proven efficacy in the treatment of inflammatory conditions of epithelia, an even wider use of ectoine can be envisaged in the future. Potential future applications include the treatment of epithelial-derived inflammatory diseases, especially nanoparticle induced, lung inflammation [98], colitis [100], and tissue protection in ischemia [105, 165]. In addition, the effect of ectoine and related compatible solutes in preventing aggregation of amyloidogenic proteins by stabilizing the native state [15, 45, 116, 117] may serve as a promising starting point for the development of amyloid-­ inhibiting novel compounds for the treatment of neurodegenerative disorders such as ­Alzheimer’s, Parkinson’s, or Huntington’s as well as prion-related diseases [16, 17, 46, 104, 166, 167]. Acknowledgments: We are grateful to the bitop AG (Dortmund, Germany) for valuable information on the industrial production and application of ectoine. Some of the authors’ work presented here has been supported by the Deutsche Forschungsgemeinschaft (DFG) and by grants from the BMBF.

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[156] Anzali S, von Heydebreck A, Herget T. Elucidation of the anti-aging effects of ectoine using cDNA microarray analysis and signaling pathway evaluation. Int J Cosmet Sci. 2009;12:381–5. [157] Buenger J, Driller H. An effective natural substance to prevent UVA-induced premature photoaging. Skin Pharmacol Physiol. 2004;17. [158] Grether-Beck S, Timmer A, Felsner I, Brenden H, Brammertz D, Krutmann J. Ultraviolet A-induced signaling involves a ceramide-mediated autocrine loop leading to ceramide de novo synthesis. J Invest Dermatol. 2005;125:545–53. [159] Pfluecker F, Buenger J, Hitzel S, et al. Complete photo protection – going beyond visible endpoints. SÖFW J. 2005;131:20–30. [160] Botta C, Di Giorgio C, Sabatier AS, De Meo M. Genotoxicity of visible light (400–800 nm) and photoprotection assessment of ectoin, L-ergothioneine and mannitol and four sunscreens. J Photochem Photobiol B. 2008;91:24–34. [161] Heinrich U, Garbe B, Tronnier H. In vivo assessment of ectoin: a randomized, vehiclecontrolled clinical trial. Skin Pharmacol Physiol. 2007;20:211–8. [162] Unfried K, Krämer U, Sydlik U, et al. Reduction of neutrophilic lung inflammation by inhalation of the compatible solute ectoine: a randomized trial with elderly individuals. Int J Chron Obstruct Pulmon Dis. 2016;11:2573–83. [163] Salapatek AM, Bates M, Bilstein A, Patel D. Ectoin, a novel, non-drug, extremophile-based device, relieves allergic rhinoconjunctivitis symptoms in patients in an environmental exposure chamber model. J Allergy Clin Immunol. 2011;127:AB202. [164] Eichel A, Wittig J, Shah-Hosseini K, Mosges RM. A prospective, controlled study of SNS01 (ectoine nasal spray) compared to BNO-101 (phytotherapeutic dragees) in patients with acute rhinosinusitis. Curr Med Res Opin. 2013. [165] Pech T, Ohsawa I, Praktiknjo M, et al. A natural tetrahydropyrimidine, ectoine, ameliorates ischemia reperfusion injury after intestinal transplantation in rats. Pathobiology. 2013;80:102–10. [166] Yang DS, Yip CM, Huang TH, Chakrabartty A, Fraser PE. Manipulating the amyloid-beta aggregation pathway with chemical chaperones. J Biol Chem. 1999;274:32970–4. [167] Ignatova Z, Gierasch LM. Inhibition of protein aggregation in vitro and in vivo by a natural osmoprotectant. Proc Natl Acad Sci U S A. 2006;103:13357–61.

Shin Haruta

6 Thermophilic photosynthesis-based microbial communities – energy production and conversion 6.1 Introduction Sunlight is the greatest and sole energy source from which our planet is continuously able to receive. Photosynthesis converts light energy from the Sun to chemical energy and supports living organisms on Earth. Photosynthetic organisms work as major primary producers in our ecosystems. The appearance and evolution of photosynthetic organisms on Earth have drastically changed the environments and afforded a variety of habitats to enhance species diversification. Photosynthesis by microorganisms contributes considerably to our ecosystems; approximately half of the primary production on Earth is achieved by microbial photosynthesis [1]. Photosynthetic microorganisms utilize chemical energy obtained through photosynthesis not only for carbon fixation but also for production of chemicals useful as fertilizers, food additives, fuels, building blocks of biopolymers, and so on [2]. So far, we have been trying to apply the abilities of photosynthetic microorganisms to various industries to develop a sustainable society. The application of photosynthetic microorganisms is widening to the fields of wastewater treatment, bioremediation, and medicine [3–5]. Light energy is abundantly supplied from the Sun, but efficient utilization in a limited space is the critical issue for biotechnological application of photosynthetic microorganisms. In order to capture light effectively, several types of photo-bioreactors have been developed: tubular type, flat panel type, vertical column, and so on [6]. Illumination by sunlight easily increases the temperature of culture solutions. High temperature is a limiting factor for utilization of mesophilic microbes, including most eukaryotic microalgae. However, a variety of photosynthetic prokaryotes that can grow over 55°C have been reported and available [7, 8]. Furthermore, operation at high temperature has the advantage of decreasing contamination and increasing reaction rates. This chapter summarizes the diversity of thermophilic photosynthetic bacteria and introduces potential applications of thermophilic microbial communities dominated by photosynthetic bacteria.

6.2 Thermophilic photosynthetic bacteria Tab. 6.1 summarizes photosynthetic bacteria. No photosynthetic archaea have been found. Cyanobacteria is the sole bacterial group harboring both photosystems, I and II, and chlorophyll and produce oxygen as a by-product. A typical example of https://doi.org/10.1515/9783110424331-006

Calvin cycle

carbon fixation pathway

Aquatic environments, soils

Soils, sediments, aquatic environments

Aerobic/ Anaerobic respiration, fermentatoin

*Chl,chlorophyll; BChl, bacteriochlorophyll **Photoautotrophic species have not been found.

Habitats

–

H2O

Electron donor

Other energy metabolisms

Sulfur compounds, H2, Organic compounds

Chl a/ β-carotene

Maojr antenna pigment

Calvin cycle

BChl a / BChl b

Chl a

Reaction center chlorophyll*

BChl a

Sulfur compounds

BChl c / BChl d / BChl e

Chlorobi

Green sulfur bacteria

Mainly hot springs

Aerobic respiration

Mainly aquatic environments

–

3-Hydroxypropionate Reductive TCA cycle/Calvin cycle cycle

Sulfur compounds, H2, Organic compounds

BChl a / BChl c / BChl d

II

Chloroflexi

Proteobacteria

Cyanobacteria

I and II

Phylum

Photosystem

Purple bacteria Green filamentous bacteria

Cyanobacteria

Tab. 6.1: Photosynthetic bacteria.

I

Soils, rice paddy soils, hot springs

Fermentatoin

–

Organic compounds

BChl g

BChl g

Firmicutes

Heliobacteria**

Hot springs

Aerobic respiration

–

Organic compounds

BChl c

BChl a, Chl a

Acidobacteria

Chloracidobacterium**

Aquatic environments, soils

Aerobic respiration (no photosynthetic growth)

–

Organic compounds

BChl a

BChl a

II

Gemmatimonadetes

Gemmatimonas Phototrophica**

154   6 Thermophilic photosynthesis-based microbial communities

6.2 Thermophilic photosynthetic bacteria 

 155

the habitats of thermophilic cyanobacteria is terrestrial hot springs. The other groups possess bacteriochlorophylls and do not utilize water as an electron source; i.e., these groups are anoxygenic. Their physiological properties and habitats are quite diverse.

6.2.1 Oxygenic photosynthetic bacteria – cyanobacteria– Cyanobacteria are widely distributed not only in aquatic environments, fresh water, and ocean but also in soils and rocks. Temperature ranges of their habitats are quite different from freezing to over 60°C. H2O is a main electron source, but some strains can also utilize sulfide similar to anoyxgenic photosynthetic bacteria (described below). Furthermore, cyanobacteria show a variety of metabolic abilities that are useful for bioremediation of pollutants and wastewater treatments [4, 5]. Cyanobacteria are traditionally classified into five groups based on their distinguishing cellular morphology: unicellular or multicellular form, nonbranching or branching filament, presence and types of differentiated cells, types of reproduction and division, etc. [9]. Thermophilic cyanobacteria, which have an optimal temperature above 45°C, are distributed in all the five groups. Molecular phylogeny based on 16S rRNA gene sequences does not correlate with the morphology-based classification. Thermophilic strains are definitely not associated with certain lineages in the molecular phylogeny. A variety of cyanobacteria retain thermophilic or thermotolerant nature. Highly thermophilic strains that are able to grow above 55°C have been reported in the following genus names [7]: Synechococcus (unicellular, both N2 fixing type and non-N2 fixing type have been known), Phormidium (non-heterocystous unbranching filament, some strains have been reported to show nitrogenase activity), Chlorogloeopsis (unbranching heterocystous filament), and Mastigocladus (branching heterocystous filament). In addition, a variety of moderately thermophilic cyanobacterial strains have been reported as Oscillatoria sp., Synechocystis sp., Gloeocapsa sp., Symploca sp., Plectonema sp., Lyngbya sp., Spirulina sp., Calothrix sp., and so on.

6.2.2 Anoxygenic photosynthetic bacteria Anoxygenic photosynthetic bacteria utilize inorganic sulfur compounds (e.g. sulfide, sulfur), H2, or other reduced inorganic compounds instead of water as electron donors for the photosynthesis (Tab. 6.1). Some also grow photoheterotrophically. Purple nonsulfur bacteria, a subgroup of the purple bacteria, show versatile metabolic abilities and their applications have been widely addressed for energy and chemical productions and bioremediation [10]. Purple sulfur bacteria, the other subgroup of the purple bacteria and green sulfur bacteria, typically utilize sulfur compounds as electron donors and grow photoautotrophically. Their sulfide oxidizing ability is useful to

156 

 6 Thermophilic photosynthesis-based microbial communities

remove a toxic and corrosive compound, hydrogen sulfide. However, these photosynthetic bacteria rarely adapt to high temperature. Not much attention has been given to biotechnological application of heliobacteria [11] and novel phototrophs in the phyla Acidobacteria [12] and Gemmatimonadetes [13]. Green filamentous bacteria, also known as filamentous anoxygenic photosynthetic bacteria, are mainly found in hot springs, e.g. the genera Chloroflexus, Roseiflexus, and Heliothrix. These grow well at 50–55°C and some can grow at approximately 70°C. Green filamentous bacteria show metabolic versatility, i.e. aerobic respiration, chemolithotrophy, and utilization of various organic compounds [14, 15]. However, few attempts have been conducted for their industrial application so far. Their thermophilic nature would be useful for various applications.

6.3 Photosynthesis-based microbial communities in terrestrial hot springs Terrestrial hot springs are promising places where solar-driven thermophilic microbial communities are obtained (Fig. 6.1). When we visit terrestrial hot springs, we are aware of colorful mat-like biofabrics in hot spring waters in the world. The biofabrics are colonies of a variety of microorganisms, so-called microbial mats. Microbial mats are well-developed microbial communities and widely occur in natural environments not only in terrestrial hot springs, e.g. sediment surfaces in fresh waters, estuaries, marine waters, hypersaline waters, and hydrothermal vents.

Fig. 6.1: Photo-image of hot spring microbial mats developed on a sediment-control dam wall at Nakabusa hot spring in Japan. Nakabusa hot spring, which is a sulfidic and slightly alkaline geothermal spring, has been attracting geochemists and microbiologists. The outflow of hot spring water emerges from seams in a sediment-control dam wall, and heterogeneous environments ranging from 75 to 50°C are observed. Several types of microbial mats develop on this dam wall: white to gray colored mats dominated by chemolithotrophic sulfur oxidizing bacteria at 75–70°C; olive green colored mats dominated by green filamentous bacteria at 69–63°C; and green colored mats dominated by cyanobacteria at 62–50°C (a piece of the mats is shown in the inset) [16].

6.3 Photosynthesis-based microbial communities in terrestrial hot springs 

 157

Microbial mats have a lot of potential applications. The densely packed forms of microorganisms are effective for the utilization of microorganisms in terms of handling, stability, and activity [17]. The proximity of microbial cells in the mats enhances interspecies interactions to stimulate community functions. As a simple example in bioremediation, oxygen production from cyanobacteria promoted the degradation activity of aerobic heterotrophs within microbial mats [18].

6.3.1 Bacterial and archaeal compositions of hot spring microbial mats Cyanobacteria-dominated microbial mats are widely distributed under hot spring waters at temperature below 60°C. Cyanobacterial mats have layered structures and grow sometimes over 3 cm in thickness (Fig. 6.1). At over 60°C, anoxygenic photosynthetic bacteria such as Chloroflexus-dominated mats are observed. In these hot spring microbial mats, photosynthetic bacteria coexist with a variety of aerobic and anaerobic bacteria and archaea, e.g. aerobic chemoorganotrophs, aerobic chemolithotrophs (such as hydrogen oxidizer), fermentative bacteria, anaerobic respiring bacteria (such as sulfate reducing bacteria and sulfur disproportionating bacteria), and methanogenic archaea. In sulfidic hot springs, sulfur-oxidizing chemoautotrophs also work as primary producers. Nitrogen-fixing ability has been found not only in cyanobacteria but also in other members of hot spring microbial mats [19].

6.3.2 Layered structure A layered structure is a notable feature of microbial mats and enables coexistence of a variety of microbes in the microbial ecosystems. For example, oxygen consumers cover the Chloroflexus-dominated mats and make anaerobic parts within the mats [20]. Even if a microbial community contained antibiotics-producing members, a layered structure would allow antibiotics-sensitive bacteria to survive [21]. Similarly, oxygenic photosynthetic bacteria can stably coexist with oxygen-sensitive anaerobes. The combination of aerobic and anaerobic metabolisms is likely effective to maintain microbial ecosystems; e.g., fermentative products such as acetic acid by anaerobic fermenters inhibit bacterial growth, but aerobic respiring bacteria immediately convert them to CO2 with consumption of oxygen [22]. Simultaneous removal of suppressive chemicals, i.e. acetic acid and O2, symbiotically supports two microbes. On the other hand, a layered structure may suppress the penetration of light and distribution of nutrients. As summarized in Tab. 6.2, photosynthetic bacteria capture a wide range of light. If bacteria in the top layer absorb certain wavelengths of light, bacteria in the lower layers utilize light of other wavelengths [23, 24]. Distribution of nutrients within photosynthesis-mediated microbial mats should change by diel cycle. However, bacteria with cellular motility may be able to move adaptively within the mats [25, 26].

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 6 Thermophilic photosynthesis-based microbial communities

Tab. 6.2: Summary of photosynthetic pigments. Wavelength (nm)

350

400

450

500

550

Chlorophylls Chlorophyll a Chlorophyll b Chlorophyll c Chlorophyll d Chlorophyll f DV Chlorophyll a DV Chlorophyll b Bacteriochlorophylls Bacteriochlorophyll a Bacteriochlorophyll b Bacteriochlorophyll c Bacteriochlorophyll d Bacteriochlorophyll e Bacteriochlorophyll g Phycobilins Phycocyanobilin Phycoerythrobilin Phycourobilin Phycobiliviolin (Phycoviolobilin) Bilin 618 Bilin 584 Carotenoids β-carotene α-carotene γ-carotene Zeaxanthin Spirilloxanthin Echinenone Spheroidene Spheroidenone Rhodopin Rhodopinal Neurosporene Black, in vitro absorption; gray, in vivo absorption.

600

650

700

750

800

850

900

6.3 Photosynthesis-based microbial communities in terrestrial hot springs 

 159

6.3.3 Productivity What are the possible products obtained from photosynthesis-based microbial communities? Gaseous compounds would be practical candidates, such as molecular hydrogen (H2) and methane (CH4), which can be continuously and efficiently collected from thermophilic microbial mats. In recent years, great attention has been paid to production and utilization of H2 as a clean energy. Among the biological hydrogen production processes that have been known [27], photosynthetic nitrogen fixation is the most possible pathway of H2 production in microbial mats. H2 is produced as a by-product of nitrogenase activity. Thermophilic nitrogen-fixing cyanobacteria have been identified as described above. Fermentative metabolism is also an important source of H2. Additionally, biophotolysis may be involved in H2 production in mats. The dynamics of molecular hydrogen within microbial mats have been studied under laboratory and natural conditions [28]. As reported in these studies, however, H2 is a useful electron donor and can be easily consumed by hydrogenotrophic prokaryotes in mats, such as aerobic H2 oxidizers, anoxygenic photosynthetic bacteria, sulfate/sulfur-reducing bacteria, and methanogenic archaea. Efficient recovery of H2 could be achieved by removing these members or disturbing interspecies hydrogen transfer [29]. Methane is a greenhouse gas, and biological methanogenesis in nature has been vigorously studied to reduce the amount in the Earth’s atmosphere. However, methane is the major component of natural gas and an attractive energy source. Methanogenic bioreactors have been operated to treat organic wastes and to recover energy. Methane-producing reactions are a part of the anaerobic respiring process of methanogenic archaea, and biological methanogenesis occurs in anoxic environments. From cyanobacteria-dominated microbial mats, methanogens have been detected, indicating anoxic area exists in the mats [30]. Fig. 6.2 shows a possible metabolic flow in solar-driven microbial communities. Fixed carbons produced by photosynthetic bacteria are converted to methane as a final product through several microbial metabolisms. In addition, methanogens also utilize H2 to produce methane directly from carbon dioxide. Since the solubility of methane in aqueous solution is quite low especially at high temperature and ambient atmospheric pressure, methanotrophs hardly exploit methane within thermophilic microbial communities. Oxygen produced through photosynthesis is removed by microbial respiration to make a part of the mats anaerobic. Oxygen respiration consumes organic compounds and H2, leading to a reduction in methane-producing efficiency. However, oxygen is readily emitted from the mats under high-temperature conditions and the layered structure helps to segregate the anoxic area from the oxygen-producing surface where the cyanobacterial population is high. Even if we do not manipulate the layered structure, methanogenesis probably occurs since methane production was observed in oxygenated water column of a lake [31]. Although oxygen respiration may slightly

160  Light

 6 Thermophilic photosynthesis-based microbial communities

CO2 O2 Oxygenic phototroph

Aerobe H2O

Saccharides

H2

Fermenter

Light

Saccharides

Methane

Methanogen

Fatty acids

SO42– Anoxygenic phototroph

SRB

CO2

H2S

CO2 Fig. 6.2: Model of material flow in photosynthesis-based hot spring microbial communities. Oxygenic and anoxygenic photoautotrophic bacteria fix carbon dioxide by utilizing water and sulfur compounds as electron sources, respectively. Organic carbons are converted through the multi-step process to methane. Molecular hydrogen produced in the communities is also utilized to convert carbon dioxide to methane. SRB, sulfate-reducing bacteria.

reduce the productivity of methane, diverse metabolic pathways would make the microbial ecosystems robust and resilient; e.g., unexpected accumulation of inhibitory substances can be removed by aerobic degradation.

6.4 Future directions – microflora engineering This chapter focused on H2 and CH4 production as the abilities of microbial communities. Photosynthetic thermophilic microbial communities have a lot of potential to produce a variety of materials if we design appropriate combinations of microbes.

6.4 Future directions – microflora engineering 

 161

It may be possible to more efficiently obtain most compounds produced in industrial fermentations, including chemical compounds, enzymes, and other proteins, through carbon fixation at high temperature. As well as lipid extraction as a biofuel from cyanobacteria [32], recovery of bacterial cellulose could be promptly realized since some thermophiles in hot springs have been known to produce cellulose [33, 34]. Application of bacterial cellulose is of interest in medical, environmental, and industrial fields. We have utilized microorganisms as a multispecies mixture for food fermentations, wastewater treatments, composting, and so on [35, 36]. Microbial communities composed of multiple species of microbes show multiple functions and sometimes higher and novel activities which cannot be achieved by using a single organism. Traditionally, we employ enrichment cultivation techniques to obtain microbial communities with desired functions from nature. In these processes, desired members are promoted, undesired members are subtracted or repressed, and some functional members are combined [e.g. 37–39]. In addition, spatial distribution is developed by biofilm or granule formation for controlling compartmentalization and proximity of the members [e.g. 40]. In microflora engineering, synthetic (microbial) ecology is the crucial discipline to design, modify, and control microbial communities [41], as synthetic biology and systems biology have made a great contribution to the biological engineering of cells. Synthetic ecology requires much deeper knowledge of how microbial communities are developed and function [42]. High throughput omics techniques advance the growth of this distinct discipline. Mathematical modeling of metabolic relationships among members in microbial communities should help to establish synthetic ecology [43–45]. Nowadays, we are surprised by findings of a variety of species-species or cell-cell interactions different from simple metabolic relationships in the prokaryotic world; competition, syntrophy, growth suppression and promotion, prey-predator, escaping and attracting, and so on. The promoting effect of heterotrophic bacteria on cyanobacteria is also highly focused on in the biotechnological field [3]. Furthermore, coexistence of other microbes would also increase the tolerance against environmental stresses [e.g. 46]. To date, bacteria and archaea in a wide range of phylogenetic lineage have been known to utilize or sense light. In these microbes other than the bacteria shown in Tab. 6.1, light energy is not enough to support their growth but is helpful to maintain viability [47, 48]. The application of illumination would also be effective for non-photosynthetic members, resulting in an increase in the functional stability of microbial communities. Some microbes sense light intensity or specific wavelength to change their metabolisms [49], although detailed mechanisms remain to be clarified. It may be possible to utilize illumination for artificial regulation of microbial community functions in the future. Design and control of microbial communities utilizing light and its energy will hopefully help and support our life in the solar system.

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References [1] Madsen EL. Environmental microbiology: from genomes to biogeochemistry. Wiley-Blackwell; Malden, MA, USA; 2008. [2] Hallenbeck PC, editor. Modern topics in the phototrophic prokaryotes: environmental and applied aspect. Cham, Switzerland: Springer International Publishing; 2017. [3] Gupta V, Ratha SK, Sood A, Chaudhary V, Prasanna R. New insights into the biodiversity and applications of cyanobacteria (blue-green algae) – prospects and challenges. Algal Res. 2013;2:79–97. [4] El-Bestawy EA, El-Salam AZA, Mansy AEH. Potential use of environmental cyanobacterial species in bioremediation of lindane-contaminated effluents. Int Biodeterior Biodegradation. 2007;59:180–92. [5] Martins J, Peixe L, Vasconcelos V. Unraveling cyanobacteria ecology in wastewater treatment. Microb Ecol. 2011;62:241–56. [6] Gupta PL, Lee SM, Choi HJ. A mini review: photobioreactors for large scale algal cultivation. World J Microbiol Biotechnol. 2015;31:1409–17. [7] Whitton BA, editor. Ecology of cyanobacteria II – their diversity in space and time. Dordrecht, the Netherlands: Springer; 2012. [8] Madigan MT. Thermophilic anoxygenic phototrophs – diversity and ecology. In: Reysenbach AL, Voytek M, Mancinelli R, editors. Thermophiles – biodiversity, ecology, and evolution. New York, NY: Kluwer Academic/Plenum Publishers; 2001. p. 103–123. [9] Casteholz RW. Phylum BX. Cyanobacteria. In: Garrity G, Boone DR, Castenholz RW, editors. Bergey’s manual of systematic bacteriology. Vol. 1. New York, NY: Springer; 2001. p. 473–599. [10] Frigaard NU. Biotechnology of anoyxgenic phototrophic bacteria. In: Scheper Th, Belkin S, Bley Th, et al., editors. Advances in Biochemical Engineering/Biotechnology. Berlin, Heidelberg, Germany: Springer; 2016;156:139–154. [11] Sattley WM, Madigan MT. The family Heliobacteriaceae. In: Rosenberg E, DeLong EF, Lory S, Stackebrandt E, Thompson F, editors. The prokaryotes; Firmicutes and Tenericutes. 4th ed. New York, NY: Springer; 2014. p. 185–96. [12] Tank M, Bryant DA. Chloracidobacterium thermophilum gen. nov., sp. nov.: an anoxygenic microaerophilic chlorophotoheterotrophic acidobacterium. Int J Syst Evol Microbiol. 2015;65:1426–30. [13] Zeng Y, Selyanin V, Lukeš M, et al. Characterization of the microaerophilic, bacteriochlorophyll a-containing bacterium Gemmatimonas phototrophica sp. nov., and emended descriptions of the genus Gemmatimonas and Gemmatimonas aurantiaca. Int J Syst Evol Microbiol. 2015;65:2410–9. [14] Hanada S. The phylum Chloroflexi, the family Chloroflexaceae, and the related phototrophic families Oscillochloridaceae and Roseiflexaceae. In: Rosenberg E, DeLong EF, Lory S, Stackebrandt E, Thompson F, editors. The prokaryotes: other major lineages of bacteria and archaea. 4th ed. New York, NY: Springer; 2014. p. 515–32. [15] Kawai S, Nishihara A, Matsuura K, Haruta S. Hydrogen-dependent autotrophic growth in phototrophic and chemolithotrophic cultures of thermophilic bacteria, Chloroflexus aggregans and Chloroflexus aurantiacus, isolated from Nakabusa hot springs. FEMS Microbiol Lett. 2019;366:fnz122. [16] Everroad CR, Otaki H, Matsuura K, Haruta S. Diversification of bacterial community composition along a temperature gradient at a thermal spring. Microbes Environ. 2012;27:374–81. [17] Bender J, Phillips P. Microbial mats for multiple applications in aquaculture and bioremediation. Bioresour Technol. 2004;94:229–38. [18] Cohen Y. Bioremediation of oil by marine microbial mats. Int Microbiol. 2002;5:189–93.

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[39] Narisawa N, Haruta S, Cui ZJ, Ishii M, Igarashi Y. Effect of adding cellulolytic bacterium on stable cellulose-degrading microbial community. J Biosci Bioeng. 2007;104:432–34. [40] Zheng D, Angenent LT, Raskin L. Monitoring granule formation in anaerobic upflow bioreactor using oligonucleotide hybridization probes. Biotechnol Bioeng. 2006;94:458–72. [41] Haruta S, Yamamoto K. Model microbial consortia as tools for understanding complex microbial communities. Curr Genomics. 2018;19:723–33. [42] Fredrickson JK. Ecological communities by design. Science. 2015;348:1425–7. [43] Mee MT, Wang HH. Engineering ecosystems and synthetic ecologies. Mol Biosyst. 2012;8:2470–83. [44] Haruta S, Yoshida T, Aoi Y, Kaneko K, Futamata H. Challenges for complex microbial ecosystems: combination of experimental approaches with mathematical modeling. Microbes Environ. 2013;28:285–94. [45] Haruta S, Saito Y, Futamata H. Editorial: development of microbial ecological theory: stability, plasticity and evolution of microbial ecosystems. Front Microbiol. 2016;7:2069. [46] Beck AE, Bernstein HC, Carlson RP. Stoichiometric network analysis of cyanobacteria acclimation to photosynthesis-associated stresses identifies heterotrophic niches. Processes. 2017;5:32. [47] Koblizek M. Ecology of aerobic anoxygenic phototrophs in aquatic environments. FEMS Microbiol Rev. 2015;39:854–70. [48] DeLong EF, Beja O. The light-driven proton pump proteorhodopsin enhances bacterial survival during tough times. PloS Biol. 2010;8:e1000359. [49] Gomelsky M, Hoff WD. Light helps bacteria make important lifestyle decisions. Trends Microbiol. 2011;19:441–8.

Lorenza Ferro, Fernanda H. B. De Miranda Vasconcelos, Francesco G. Gentili and Christiane Funk

7 Photosynthesis at high latitudes – adaptation of photosynthetic microorganisms to Nordic climates 7.1 Introduction 7.1.1 Photosynthetic microorganisms: characters of a billion years-old story Microalgae and cyanobacteria are photosynthetic microorganisms responsible for approximately half of the oxygen production on Earth, being the primary producers in oceans and seas. Naturally acting as “solar cell factories,” these microorganisms can convert sunlight, CO2, and few other nutrients into energy and biomass. Despite their small size, in the range of a few microns, algae are ubiquitous: as aquatic organisms, they are usually found in marine as well as freshwater environments, where they live suspended in water or attached to sediments, plants, and animals. Some species can even colonize terrestrial environments, rocks, ice, and snow [1]. Looking at their biodiversity, numbers are quite impressive: up to 1 million species are estimated to live on Earth. Interestingly, many of them are still unknown or not characterized yet [2, 3]. Cyanobacteria, also known as “blue-green algae,” are photosynthetic p ­ rokaryotes; their evolutionary appearance about 3.5 billion years ago significantly contributed to a change of the atmospheric composition on our planet. Being similar to other bacteria in size, morphology, inner structure, and biochemistry, cyanobacteria additionally possess pigments and proteins imbedded in special biological membranes to perform oxygenic photosynthesis. Cyanobacteria can exist in different forms: unicellular, filamentous, planktonic or benthic, and coccoid (Fig. 7.1).

Fig. 7.1: Filamentous (a) and coccoid, colony-forming (b) cyanobacteria isolated from Lake Nydala, Umeå, Sweden (1000× magnification). (Photos: Lorenza Ferro) https://doi.org/10.1515/9783110424331-007

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They are found almost everywhere, but particularly in lakes and oceans, where, under certain conditions, they reproduce exponentially to form blooms. Most of them are innocuous, but some can produce cyanotoxins; during blooms, these toxins reach high concentrations and can poison or even kill animals [1]. Some cyanobacteria even can fix atmospheric nitrogen; ironically, nitrogenase, the enzyme being responsible for the reduction of N2, is extremely sensitive to oxygen. Nitrogen fixation therefore either must be temporal or spatial separated from oxygenic photosynthesis. Some filamentous cyanobacteria differentiate special cells called heterocysts, which lack the oxygenic photosystems and possess a glycolipid cell wall keeping the oxygen concentration sufficiently low for nitrogen fixation [4]. Microalgae are eukaryotic cells, which may have appeared on our planet about 2 billion years ago. According to the “endosymbiotic theory,” eukaryotic algae evolved when a nonphotosynthetic eukaryote engulfed a cyanobacterium and, instead of digesting it, adopted it. Over time, this cyanobacterium became an integrated organelle, the chloroplast [5]; genes were transferred from the cyanobacterial genome to the algal nucleus. Besides nucleus and chloroplast, algal cells further contain mitochondria, Golgi body, endoplasmic reticulum, and other typical eukaryotic organelles. Similar to cyanobacteria, also microalgae are found in different shapes: amoeboid, capsoid, coccoid, filamentous, flagellate, and sarcinoid (Fig. 7.2). According to their endosymbionts, structural diversity, biochemical pathways, reproduction, and ecology microalgae are classified in several supergroups (Fig. 7.3). Between these, the most well known are green and red algae, dinoflagellates, cryptophytes, euglenoids, and the most diverse, heterokont algae, which include the ubiquitous diatoms [1, 6]. Through photosynthesis, plants and cyanobacteria fix carbon dioxide from the atmosphere into organic substances, for example sugar, starch, cellulose, and other useful molecules. The energy used for this process is supplied by sunlight. Because photosynthesis is based on carbon dioxide, plants absorb this molecule produced by industrial processes and recycle it; photosynthetic microorganisms therefore actively

Fig. 7.2: Green microalgae (a) Coelastrum astroideum and (b) Scenedesmus sp. found in Swedish freshwater and wastewater streams (400× magnification). (Photos: Lorenza Ferro).

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Cyanobacterium Primary endosymbiosis

First photosynthetic eukaryote

Glaucophytes

Green algae Plants

Red algae

Secondary endosymbiosis(es)

Chromophytes: Heterokonts, Haptophytes, Cryptophytes, Dinoflagellates

Fig. 7.3: Schematic view of the evolution of some phylogenetic algal supergroups.

contradict the greenhouse effect and global warming. At the same, time algal and cyanobacterial cells produce a wide range of different biomolecules [7–9]. Carbohydrates are produced and stored mainly as cellulose in the cell wall and starch (or starch-like products) in the plastids. Algal carbohydrates can be easily converted into fermentable sugars for biofuel production due to the negligible content of hemicelluloses and lignin [10]. Proteins can reach more than 60% of biomass dry weight and are mainly produced by algae during the exponential growth phase. They represent an essential source of nitrogen in case of depletion in the environment. Proteins from algae recently gained interest for food and feed applications, due to their high nutritious quality; they even are superior to conventional plant proteins [11–13]. Lipids are mostly accumulated during the stationary phase of algal growth and can approach 20–60% of dry weight, depending on algal species and growth conditions. They are classified as storage lipids (nonpolar), mainly in the form of triacyglycerides (TAGs), and structural lipids (polar), like fatty acids (especially polyunsaturated fatty acids, or PUFAs), phospholipids, wax esters, and sterols. TAG molecules can be converted into biodiesel through a process called transesterification [14–16]. The omega-3 long-chain PUFAs, like eicosapentaenoic acid (EPA, 20:5 n-3) and docosahexaenoic acid (DHA, 22:6 n-3) are recognized as beneficial for human health and can be supplemented in nutrition and medicine [17–19]. Algae can also produce many other high-value compounds of commercial interest, e.g. pigments, vitamins, antioxidants, and molecules with antifungal, antimicrobial, and antiviral properties [7, 20–23].

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7.1.2 How, what, where? Growing algae in northern environments During the last decades, microalgae raised great interest as a source of biomass for biofuels, green chemicals, bioplastics, pharmaceuticals, food, and feed, due to their unique and diverse biochemical composition and their ability to grow faster and at higher densities than land crops, with no agricultural land requirements [24]. Even more, some microalgal species are able to grow on wastewaters and polluted environments and can be successfully used for bioremediation [25]. Despite this great potential and the increasing research in the field, microalgal cultivation for biomass production at commercial scale still remains a challenge [26–28]. Each step in the algal production chain presents aspects that need to be carefully evaluated and improved, in order to obtain the highest efficient process at lowest cost. Not only engineering- and industrial-related issues (e.g. bioreactors/ponds design, biomass harvesting and drying, extraction, refining) and energy balances but also – and above all – fundamental factors like light, temperature, pH, and nutrients, which effectively influence algal growth and biomass quality, have to be analyzed. As mentioned before, there are hundreds of thousands of algal species, showing very different physiological response to specific environmental conditions. The choice of the algal species for any particular process (i.e. the best microorganism with the highest/fastest biomass productivity and/or with the highest content of a valuable compound) is then of extreme importance: collection, screening, and characterization of algae are therefore crucial preliminary steps for the whole production system, for its feasibility, sustainability, and efficiency. Moreover, to avoid additional energy inputs, to minimize the costs and to make the most of natural resources, local algae, well adapted to the specific climate in which the process will be established, should be chosen. Before asking “how” to grow algae and “how” to optimize the downstream processes, one should first analyze “what” to grow and “where” to grow it. This is particularly important in areas characterized by relative extreme climate conditions, like the northern countries. In these environments, great variation in sunlight availability during summer and winter, as well as low temperatures, has a strong impact on photosynthetic microbial communities. In this context, the Swedish consortium “MicroBioRefine,” presented at the end of this chapter, is an excellent example of the development of a local-sustainable bioprocess in which characterization of (new) photosynthetic microorganisms from northern habitats allows the exploitation of natural resources in a specific (and unfavorable) environmental context. As presented above, light and temperature are among the main factors influencing algal growth, biomass productivity, and its biochemical composition, making algal parks challenging in northern countries: i.e. during the dark autumn/winter months, photosynthetic light-harvesting is challenging, but at the same time, the bright sunlight available from March to September might damage the photosynthetic organisms. At the same time, the organisms must adapt to a wide range of temperatures: cold temperature resistance is important at the end of the winter when high

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light intensity is coupled to low atmospheric temperature. In the following sections, we will discuss how photosynthetic microorganisms successfully survive in the extreme conditions of light and temperature at high latitudes, with reference to their adaptation strategies at biochemical and molecular level (Tab. 7.1).

7.2 Acclimation strategies of photosynthetic microorganisms living at high latitudes 7.2.1 Light acclimation Light is the energy source that drives photosynthesis. Biochemically, photosynthesis can be divided into two stages: the light-dependent “light reaction” and the light-independent “dark reaction.” The light reaction occurs in two pigment-binding protein complexes, called Photosystem I (PSI) and Photosystem II (PSII), imbedded in a special system of biological membranes, the thylakoid membranes. The set of pigment molecules involved in the evolution of one molecule of O2 are defined as photosynthetic units (PSUs). Both photosystems are connected by a protein complex called cytochrome b6/f. Sunlight is captured by the pigments of the light-harvesting complexes surrounding the photosystems and is transferred to the reaction center, where the photochemical reaction takes part. Two electrons, extracted from a molecule of water, are transferred from PSII via cytochrome b6/f to PSI, using the light energy to produce a molecule of nicotinamide adenine dinucleotide phosphate hydrogen (NADPH2). At the same time, protons are translocated into the intrathylakoid space, the lumen, to form a pH gradient, which in turn drives adenosine triphosphate (ATP) synthesis through a membrane-bound enzyme called ATP synthase. Both ­biochemical Tab. 7.1: Main acclimation strategies put in place by photosynthetic microorganisms when exposed to conditions of low/high light intensity and low temperature in northern climates. LOW LIGHT

HIGH LIGHT

LOW TEMPERATURE

–– Increase of chlorophyll pigmentation –– Decrease of photoprotective pigments –– Heterotrophy or mixotrophy –– Dormancy induction, cysts and spores formation –– Reduced metabolism and cell division –– Quantum coeherence

–– Photoinhibition and heat dissipation (short-term) –– Change in PSI/PSII ratio (cyanobacteria) –– Xantophyll-cycle (microalgae) –– UVR-damage repair mechanisms of DNA and proteins –– Antioxidant molecules and enzymes –– Gliding motility –– Photoprotective compounds

–– Increase of membrane fluidity and stability –– Production of intra- and extra-cellular cold-protective compounds –– Cold-adapted proteins and enzymes –– Reduction of PSII size, increase of carotenoid/chlorophyll a ratio –– Increase of photosynthetic energy sink capacity, photostasis maintenance

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reductant and biochemical energy (NADPH2 and ATP, respectively) will then be used in the dark reaction to fix inorganic carbon, which is performed by the enzyme ribulase-bisphosphate-carboxylase/oxygenase (Rubisco) [29, 30]. During the fixation process, the substrate CO2 (which decreases during photosynthesis) competes for the active site of Rubisco with O2 (which increases during photosynthesis). Therefore, both cyanobacteria and algae have developed carbon concentrating mechanisms that allow them to concentrate carbon at levels higher than the ambient and keep the supply of CO2 sufficient [31, 32]. Furthermore, algae are able to take up several different sources of organic carbon [33], which burst the algal production under limiting conditions of resources. The carbon is incorporated in the cells together with phosphorus (phosphate PO4) and nitrogen (ammonium NH4 and nitrate NO3) to build up complex organic molecules. Only a small part of the solar radiation, ranging from 400 to 700 nm of the electromagnetic spectrum and denoted as PAR (photosynthetically active radiation), can be used in photosynthesis [29, 34]. The photosynthetic activity of higher plants and algae is classically described by a light-response curve (P/I), which shows the effect of irradiance on photosynthesis. At low irradiance, the photosynthesis rate depends linearly on the light intensity (α, or quantum yield [QY]), but when light increases, this dependency is gradually lost and photosynthesis becomes less and less efficient, until it reaches a plateau called maximum rate of photosynthesis (Pmax). With further increases in the irradiance, photosynthetic rates decline from their light saturated value, a phenomenon called photoinhibition, which in the worst case leads to damage of the photosystems or the whole organism [35]. While at high latitudes (>50°), light availability averaged over the year is extremely reduced, during the short summer period (usually less than 3 months per year), solar irradiance can reach very high levels, potentially higher than on any other location [36]. Due to the drastic changes in light quantity and quality faced in the Nordic habitats over the year, photosynthetic organisms had to develop different photoacclimation strategies to adapt and survive. Nearly continuous light during the summer months are followed by extremely short daily photoperiods during autumn and winter. In addition, seas, lakes, and rivers in these regions often are completely frozen for several months, resulting in a further reduction of solar radiation passing through the ice and snow at water surface. Plants adapted to these changes with various long-term and rapid adaptation strategies. Long-term acclimation occurring seasonally involves structural, morphological, and biochemical changes within the chloroplast and the plant cell. To rapid illumination changes, plants react through mechanisms of state transitions, coupling the photosynthetic light harvesting complex from one photosystem to the other, fluorescence, or heat dissipation to release excess light energy in a harmless way performed with the help of carotenoids. Instead of using light energy in the photosynthesis, these process protect from too much light absorption and therefore also are called nonphotochemical quenching [37–42]. Low and high light acclimation strategies will be discussed separately in the following sections.

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Low light Plants acclimate to long-term low irradiance by increasing either the size or the number of PSUs. The increased amount of light-harvesting pigments, i.e. chlorophyll a, allows more light energy to be absorbed and transferred to the reaction centers, and the PSUs become more efficient. At the same time, when the number of PSUs per cell is increased, Pmax increases and more light is then required to reach s­ aturation. Increased chlorophyll pigmentation is accompanied by a general decrease in photoprotective pigments, like carotenoids, to minimize the dissipation of light [34, 43–46]. These mechanisms have been found in all known algae; however, unusually high chlorophyll a concentrations have been reported from diatoms, which are the dominant microalgal species in Polar regions. Diatoms also produce chlorophyll c as an accessory pigment, which enables them to more efficiently use the available light radiation, even below ice [47, 48]. Certain classes of algae are extremely well adapted to low-light conditions, ­allowing them to grow deep in the water column or survive in the Nordic climate. Cryptophytes, ubiquitous unicellular species evolving after two endosymbiotic events (Fig. 7.3), contain (besides the membrane-integrated light harvesting antenna) a soluble antenna consisting of phycobiliproteins (PBPs) binding the pigment phycoerythrin. Also cyanobacteria and red algae contain these extrinsic light harvesting apparatus; however, even though the PBPs originated from the red algal ancestor, in cryptophytes, they are not organized into phycobilisomes bound to the stromal face of the thylakoids, but are located in the thylakoid lumen [49, 50]. Light energy is then driven to the reaction centers and will be converted into chemical energy. Recently, it was observed that the energy transfer within the PBPs is not random, instead the fastest possible route is taken. This mechanism, known as “quantum coherence,” is controlled by the quaternary structure of PBPs, switching from an open to a closed conformation. Quantum coherence allows improving the energy usage efficiency and, thus, survival in harsh light conditions [51]. Generally, minimal irradiance is required to support the life of phototrophic cells [52]. To survive even in total darkness, a typical situation faced by microorganisms living below snow or ice, algae had to develop special strategies [53–56]. Some microalgae can switch their metabolisms from photoautotrophy to heterotrophy (or mixotrophy); their energy production is driven by organic carbon sources instead of photosynthesis [57]. Specific active transport systems enabling the uptake of carbohydrates, including acetate, glucose, lactate, and glycerol, have been found in several green and red algae and diatoms [58, 59]. Other common strategies are the induction of a dormancy state or the formation of resistant cysts and spores. During dormancy, the metabolic rate, as well as cell division and carbon concentration, is slowed down. While the algae are able to survive with minimal requirements over the winter, the preserved energy allows them to reactivate and grow rapidly when ambient light conditions improve [60–63]. Cyst and spore formation is a mechanism prevalently used by diatoms and dinoflagellates but has also been found in some chlorophytes

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and cyanobacteria (in the latter called akinetes) [63–67]. This resting stage is characterized by different morphology and physiology compared to vegetative cells. The resistant “outer coating” of cysts and spores provides algal survival against several environmental extremes and over a long time period. Germination of these cysts may significantly contribute to algal bloom formations [63, 68]. High light As mentioned before, photosynthetic microorganisms living in cold regions need to be efficiently adapted to fluctuations of light availability, due to the strong seasonality of solar irradiance and environmental conditions (e.g. ice formation and melting). While living with low or no light for the most part of the year, these algae can experience even very extreme irradiances during the summer period. The excess of PAR generally induces a process called photoinhibition, which involves a decrease in the energy conveyed to PSII (QY) and an increase of dissipation through non-­photochemical reactions (NPQs). These important photoprotection mechanisms help to prevent/limit overexcitation of the photosystem reaction centers and therefore photodamage [34]. Contrary to sessile higher plants, many microalgae and cyanobacteria are motile due to the presence of special flagella, which drive photo- and chemotaxis, the movement in response to light and chemical stimuli, respectively [69–72]. Additionally, so-called gliding motility allows them to nonflagellar translocation mediated by solid or viscous substrates and enables them to escape from high irradiances and optimize their position in the environment. This kind of motility is typical of some pennate diatoms, red algae, and filamentous cyanobacteria [73, 74]. Light-dependent damage is nevertheless unavoidable in photosynthetic organisms; therefore, efficient repair systems need to be available, in order to rapidly replace the malfunctional components [75]. The main target of light-induced damage is the D1 protein (or PsbA), the core protein of the PSII reaction center, which has high turnover rates (high degradation and frequent replacement) [76–78]. Photodamage also is induced directly by the reduction of the plastoquinone pool (acceptor side photoinhibition), light damaging the oxygen-evolving complex of PSII (donor side photoinhibition), or the manganese ions of the oxygen-evolving complex (­manganese mechanism) [79–83]. High light further induces the formation of reactive oxygen species (ROS), with highly oxidizing and toxic effects on the photosynthetic ­apparatus. Rather than directly attacking the reaction centers, ROS have been found to inhibit the repair systems by repressing the synthesis of new proteins [80, 84]. Nonphotochemical quenching can consist of state transition quenching, the rapid energy-dependent quenching (also called feedback de-excitation) and photoinhibitory quenching. State transitions in photosynthetic organisms allow balancing the excitation energy given to each of the two photosystems in order to maintain a high photosynthetic rate. The distribution of the absorbed light energy occurs through migration of the light-harvesting complex (e.g. LHCII in green algae, phycobilisomes

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on cyanobacteria) between PSII and PSI. In plants, this phenomenon is well studied and revealed a very complex mechanism involving LCHII kinases and phosphatases, the recruitment of cytochrome b6/f, and the switching of electron flow from cycling to linear [42]. In cyanobacteria, high irradiances induce additionally a change in the ratio of PSI/PSII, the amount of the highly abundant PSI decreases in the thylakoid membranes, and at the same time the amount and size of the phycobilisomes connected to PSII diminish [85–89]. Also the soluble orange carotenoid protein has been found to be important for cyanobacterial NPQ [90]. In microalgae, a xantophyll cycle is involved in NPQ; the oxidation of a carotenoid (e.g. violaxanthin, antheraxanthin, or diadinoxanthin to zeaxanthin or diatoxanthin) promotes the dissipation of energy as heat, thus helping to prevent photoinhibition. Furthermore, special stress-induced proteins with high sequence similarity to the light-harvesting antenna of the green linage, called light-harvesting like proteins (Lil), have been found to be important for NPQ in cyanobacteria and plants [91]. Sunlight further contains harmful ultraviolet radiation (UVR), which at high doses can irreparably deteriorate the photosynthetic apparatus and induce DNA mutations. A common UVR effect found in several red algae is the alteration of the chloroplast structure, due to disruption of thylakoid membranes and displacement of pigments. UVR therefore inhibits essential cellular processes like photosynthesis and nuclear division and finally may lead to cell death. Algae, especially ice- and snow-living algae, evolved active mechanisms to protect against excessive amounts of UVR [92–96], e.g. the ability to repair and replace cellular damaged components or damaged DNA. UVR (specifically UVB) absorption induces DNA single-strand breaks, dimerization of adjacent nucleotides, or cross-links between nonadjacent nucleotides, which influence DNA transcription and translation and induce mutations with possible lethal effects. Algal cells (similar to higher plants) repair DNA photoproducts either by photoreactivation or base excision repair [97]. Photoreactivation is mediated by the enzyme photolyase, which upon light induction binds to damaged DNA and specifically removes pyrimidine dimers. Base excision repair is a light-independent process found in all eukaryotes [98, 99]: a specific enzymatic complex recognizes, cuts, and removes the damaged DNA fragment, and the gap then is filled by the enzymes polymerase and ligase. Ultraviolet (UV) light also can damage proteins [100]. As previously mentioned, the PSII reaction center protein D1 is most sensitive to light, and its turnover-mediated repair cycle seems to be connected to UVR tolerance in microalgae [101–103]. UVR can further damage proteins indirectly, since oxygen released from photosynthesis can be chemically converted to ROS upon UV induction. Algae protect themselves from ROS by producing antioxidant molecules, including vitamins, carotenoids, and reduced glutathione, or efficient antioxidant enzymes like superoxide dismutase, catalase, glutathione peroxidase, and other enzymes involved in the ascorbate-glutathione cycle [56, 104, 105]. These “passive” mechanisms of UVR protection work like sun screen: robust cell walls partially filter UV light and carotenoid pigments, absorbing above 400 nm in the visible spectrum, and completely screen off

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violet and blue light, thus providing an additional protection barrier [92, 99]. Some algae and cyanobacteria also developed UV-absorbing pigments, e.g. mycosporinelike amino acids, photochemically stable molecules that maximally absorb UVA and UVB light (between 310 and 360 nm). Dinoflagellates display the highest variation in UV-absorbing compounds, explaining their successful diffusion in UV-rich environments [99, 106].

7.2.2 Cold adaptation Psychrophiles and psychrotolerant The climate at high latitudes is determined by cold. While summer temperatures have an average of 10–15°C, in winter, temperatures are below freezing point and can reach even −40 to −50°C. The persistent cold often comes with freeze-thaw cycles, high fluctuation of solar irradiance, and great variation of nutrient availability [107]. Life in these regions is therefore extremely challenging, and living organisms need a special adaptation to survive such tough conditions. Biochemical and molecular processes in a cell require a specific temperature range, usually between 20 and 40°C. Below the optimal temperature, the cell metabolism slows down until it stops, leading to cellular damage and, eventually, to cell death [108, 109]. Despite these requirements, photosynthetic microorganisms have successfully colonized cold environments, even extreme cold ones: some algal species are found on snow (Fig. 7.4) or ice surface or melt pools, on glaciers, or in brine channels in the sub-ice platelet, where temperatures can reach −20°C [110]. In these locations, photosynthetic microorganisms can reach very high densities, representing the main primary producers and sustaining the ecology of these areas. The biodiversity of photosynthetic microorganisms living in cold environments is surprisingly high. Generally, cold living organisms are classified into “psychrophiles” and “psychrotolerant,” depending on their temperature tolerance for growth. ­Psychrophiles are organisms with an optimal temperature below 15°C, which are predicted to metabolize up to −40°C and to survive even lower temperatures. However, these “cold-obligates” will not survive temperatures higher than 20°C. Psychrotolerant algae have growth optima in the range of mesophilic organisms (20–40°C) but are able to grow at lower temperatures with diminished rates. These “cold-facultative” organisms are the most common ones found in cold environments [111]. Prokaryotes When the first cyanobacteria evolved during the Precambrium, the climate on Earth was extremely cold, with long glacial periods. The ability to adapt and survive to these harsh conditions still is found in present-day cyanobacterial species, explaining their successful ecological diversity in modern cold environments [112]. Many species of cyanobacteria have been observed in low-temperature regions, both filamentous

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(e.g. Oscillatoria, Nostoc, Phormidium) and unicellular (e.g. Synechococcus and Chroococcidiopsis). They are most common on ice shelves and in freshwater environments but can also grow within soil crusts, within rocks, or symbiotic in lichens. In contrast, they are nearly absent in cold marine environments [110, 113]. Cyanobacteria in general are not considered as true psychrophiles; their optimal growth temperatures typically range between 15 and 35°C. Their growth rate in cold regions is therefore quite low, implying that they are not genetically adapted to low temperatures. Their dominance in polar regions could be explained by competitive advantages like tolerance to desiccation, freeze-thaw cycles, low nutrient supply, and/or high solar radiation [110, 112, 114]. Eukaryotes Among psychrophilic microalgae, diatoms are most abundant and are present in marine environments, both cold seas and sea ice. Due to their excellent adaptation to cold environments and much faster growth rates than psychrotolerant cyanobacteria, diatoms are the primary producers in polar regions, where they produce large visible “blooms” in spring and are the main component of the food chain [107, 112, 115]. The most representative species are Nitzschia, Pinnularia, Navicula, Melosira, and the chain-forming Amphiprora [110]. In addition to diatoms, so-called “snow algae” colonize cold environments, conferring a pink, red, green, or yellow color to snow [116]. The best-known specie is Chlamydomonas nivalis, a green alga with red spores, which produce the reddish “watermelon snow” during spring and summer (Fig. 7.4), when snow melts and more solar radiation and nutrients become available [115, 117, 118]. Chlamydomonas raudensis is another well-characterized and

Fig. 7.4: “Watermelon snow” in Arctic landscape due to the presence of Chlamydomonas nivalis. (Source: www.earthnetwork.news).

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ecologically important green alga growing in ice-covered lakes [115]. Strain UWO241, the best studied one, will be presented in detail at the end of this paragraph. Other common microalgal species growing on or within snow and ice are Chloromonas, Chlorella, Ankistrodesmus, and Mycanthococcus [107, 115, 119]. Do or die: adaptation strategies to cold temperatures The adaptation of phototrophic microorganisms to low temperatures at high latitudes involves a wide range of mechanisms, which partially overlap with those used to adapt to darkness or low light: seasonally induced dormancy or induction of specialized enzymes and cryoprotectants to avoid cell lysis [56, 107]. Moreover, special strategies are needed in psychrophilic and psychrotolerant microalgae and cyanobacteria to balance light absorption, energy transfer, and chemical energy production (ATP and NADPH) with the downstream consumption of photosynthetic products at low temperatures [108, 120, 121]. Membrane fluidity At cold temperatures, photosynthetic microorganisms have to regulate the lipid content of their biological membranes to improve their fluidity, which allows survival as long as the cytoplasm remains unfrozen [110, 112, 122, 123]. PUFAs, a class of shortchain desaturated lipids, are produced and incorporated into the membrane [124]. The enzyme acyl-lipid desaturase, activated by cold-shock [125], desaturates lipids. The fluidity of the membranes is further increased by synthesis of polar lipids, like zeaxanthin, which stabilize the membrane compared to nonpolar lipids. Additionally, the membrane protein content is diminished, since lower protein-lipid interactions improve the flexibility of acyl chains. Even an increase in cis- and a decrease in trans-fatty acids were observed during cold acclimation [126]. At the same time, solutes as trehalose are released into the cell to lower the freezing point of the intracellular water. These processes not only improve the fluidity of the membrane but also reduce cell desiccation since the osmotic equilibrium is best retained [113]. The production of extracellular compounds, usually polymeric molecules, further helps to protect the cells and prevents freezing damage and allows algae and cyanobacteria to survive the prolonged seasonal dormancy in frozen or liquid water. Between a few minutes to some hours after thawing, the adapted microorganisms are able to fully regain their photosynthetic activity [112]. Besides containing a strong, rigid cell wall to protect from drought and freezing, the snow alga Chlamydomonas nivalis can even secrete carbohydrates in the surrounding environment, producing a mucilage sheet, which attracts bacteria and traps particles into the snow. These trapped particles help the alga to increase the absorption of sunlight, which in turn warms the snow, induces its melting to make liquid water available, and forms so-called “cryoconite holes” [127–129].

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Enzymatic adaptation The rate of any biochemical reaction is dramatically affected by a decrease in temperature: lowering the temperature by 10°C causes biochemical reaction rates to decline two to three times [120]. In cold conditions, enzymes usually are denatured, losing their native folding, flexibility, and eventually their catalytic activity. Low temperatures therefore may result in impaired metabolism, molecular structure, cellular transport, and decreased rates of DNA replication and translation. Moreover, at 0°C, the viscosity of water is strongly decreased, further impairing all biological processes [130]. Despite the critical reduction in enzymatic activity, psychrophilic microorganisms growing at low temperature show a doubling time very similar to that of mesophilic bacteria grown at 37°C [131]. Photosynthetic microorganisms living in cold environments developed specific cold-adapted enzymes characterized by an increased structure flexibility, allowing them to resist freezing-temperature-induced damages like protein aggregation and precipitation; instead, they maintain appropriate metabolic reaction. These enzymes have a high specific activity, high turnover number, low activation enthalpy, and low substrate affinity at low or moderate temperatures but are inactivated at increased temperature rates [56, 120, 130, 132, 133] As an example, the enzymes nitrate reductase and argininosuccinate lyase isolated from the psychrophilic green alga Chloromonas showed maximal activity and thermal stability at low temperatures, while the same enzymes from the mesophilic alga Chlamydomonas reinhardtii were completely inactivated at 5°C [134]. Common features characterizing proteins in cold-adapted species include a high degree in polar residues and few hydrophobic residues resulting in high interaction with solvents; additionally, they have very few hydrogen bonds and ion pairs, lack salt bridges, and have only a few subunit interactions [133, 135]. Photosynthesis and energy balance Photosynthetic microorganisms growing in cold environments are in a constant state of energy imbalance; while light absorption is temperature independent, the use of electrons and equivalents in photosynthesis (i.e. CO2) is reduced [120, 136]. Psychrophilic phototrophic organisms respond to this imbalance in different ways; photostasis is attained either by reducing the light-harvesting antenna size and/or by dissipating energy as heat, through non-photochemical quenching. Similarly to high PAR or UV stress, the size of PSII can be reduced significantly at low temperatures by increasing the carotenoid/chlorophyll a ratio [137, 138]. Alternatively, the photosynthetic sink capacity is increased by overproduction of Calvin cycle (“dark reaction”) enzymes [120]. In diatoms, an adaptive strategy to compensate for the reduced metabolism at low temperatures was observed. These algae uptake nitrate in levels exceeding the nutritional requirements to balance the photosynthetic

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energy production [139]. The diatom Thalassiosira pseudonana not only consumes excess energy via NO3− reduction but even photorespirates as complementary energy sink to respond to the energy imbalance between photochemistry and cellular metabolism [140].

7.3 The strange case of Clamydomonas raudensis strain UWO241 Defined as “unusual psychrophilic green alga,” Clamydomonas raudensis strain UWO241 was isolated from Lake Bonney, Antarctica, one of the driest and coldest regions on Earth. Lake Bonney is roughly 40 m deep and permanently covered by 3 to 4.5 m of ice [141]. Because of this thick layer of ice, vertical mixing of the water column by wind is prevented, giving rise to vertical stratification with strong temperature and salinity gradients. Temperatures range from 0/−1°C (bottom of the lake and ice-water interface respectively) to 6°C (at a depth of 14 m), while the salinity increases from freshwater at the surface to a hypersaline brine at the bottom of the lake [142, 143]. Antarctica is characterized by 4 months of total darkness during the winter season, while in summer, the ice layer covering the lake can absorb up to 99% of incident PAR, screening light with a wavelength higher than 600 nm [144]. In such conditions, photosynthetic microorganisms populating the water column need to be adapted to low light, specifically in the blue-green spectrum [145]. Biological investigations of Lake Bonney date back to the sixties and seventies, when several microbial communities were found, including cryptophytes, chrysophytes, and chlorophyte algae (from 4 to several meters of depth); oscillatorious cyanobacteria (especially at the ice-water interface); ciliates and rotifers; protozoa; bacteria; and fungi [141, 146, 147]. Two decades later, the “enigmatic” green alga designated as UWO241 was isolated from a depth of 10 to 17 m at high salinity (approximately 700 mM). In these depths, in a transition zone between an upper oxygen-rich and a deeper oxygen-deficient layer [146], the average temperature is 4–6°C and irradiance is below 15 μmol photons m−2·s−1 [145]. UWO241 is a true psychrophile, unable to grow at temperatures above 16°C [148, 149]. The biflagellate single cells have an ellipsoid or ovoid shape, being 10–15 µm long and 5–12 µm wide. Its asexual reproduction occurs through zoospore formation, each sporangia containing 16 to 32 single motile cells. Photosynthetic pigments in UWO241 are essentially the same as in all the other green algae, with lutein and neoxanthin being the most abundant carotenoids. Interestingly, the thylakoid membranes in UWO241 resemble those of psychrophilic diatoms, with their high content of galactolipids and PUFAs and the lack of grana stacking. These morphological changes promote the energy distribution between the two photosystems [120]. Moreover, special eyespots are located in-between the plasma and thylakoid membranes, which might act as like an optical system to adjust the amount of absorbed light through phototaxis [150].

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The most unusual feature of this microorganism is its photosynthetic apparatus: the amount of PSI and proteins associated with PSI is low, and so is the chlorophyll a/b ratio. Chlorophyll b is bound by the light harvesting antenna, and UWO241 therefore has enhanced light-harvesting capacity. Even the concentration of xanthophyll cycle carotenoids, which prevents energy dissipation and maximizes photosynthesis (e.g. antheraxanthin and zeaxanthin), is low [148, 151]. The light-harvesting complexes of PSII absorb mainly the energy-rich blue light; the high PSII to PSI ratio of UWO241 is therefore thought to be an advantage for growth under constant exposure to low blue light levels. The strategy of C. raudensis strain UWO241 to produce photosynthetic energy entirely using PSII, while PSI takes care of spillover energy [120], makes this psychrophilic alga able to survive in very low irradiances with narrow light quality and high ATP requirements [148, 152]. In this strain, the highest rate of oxygen evolution, which indicates maximal photosynthetic efficiency, was recorded at 8°C [150]. UWO241 is naturally low light adapted but can adapt to high irradiances. It has been found that it can easily grow at 8°C at a light intensity 15 times higher than its ambient growth irradiance [120]. Nevertheless, it lacks the state-transitions mechanism for energy distribution between the two photosystems (short-term photoacclimation; see above) [152]. Instead, its energy balance is regulated by the amount of the reaction center proteins in PSI and PSII [153]. Photoinhibition in this psychrophile alga is only dependent on light, but not on temperature; at low temperatures, full and rapid recovery of photosynthesis occurs due to the presence of a novel D1 repair cycle within PSII, which efficiently operates at cold conditions [154]. C. raudensis also exhibits an unusual growth response dependent on the light quality: it grows exponentially under white or blue light, but growth is inhibited under red light, which is lacking in its natural environment. However, cells grown under red light remain viable and resume their growth once transferred to white light [153]. To date, no complete sequence of any genome of photosynthetic psychrophiles is available. Chlamydomonas raudensis UWO241 would be a very interesting candidate, its adaptive mechanisms being studied for more than a decade. Moreover, due to the particular characteristics of its natural habitat, strain UWO241 could be used as an important indicator of environmental changes, e.g. to monitor ozone depletion, UV radiation, pollution, and climate warming in Antarctic region [120].

7.4 MicroBioRefine – biomass production and wastewater ­reclamation in Nordic climate For converting a society dependent on fossil fuels and nuclear energy into a sustainable, bio-based society, all production processes and products must be optimized for low material and energy use. In addition, new bio-based approaches need to be explored that utilize regional synergistic effects of material and energy flows. In Umeå, Northern Sweden, a consortium has been established from researchers in different

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disciplines working on three North Swedish Universities and one research institute. Aim of the work is to investigate recycling of carbon dioxide, nutrients, and pollutants from flue gases and wastewaters using photosynthetic microorganisms adapted to the Nordic climate. The biomass produced by these microorganisms is used for biorefining to produce mainly biofuels such as biodiesel, bioethanol, and biogas, but also to produce energy-rich products like animal feed and biofertilizers. Instead of providing expensive and energy-demanding fertilized media to the algae and cyanobacteria, the microorganisms are growing on sewage and flue gases and therefore participate in preserving the environment. Local strains adapted to the harsh Nordic climate and extremophiles from culture collections are compared to well-studied microorganisms grown in open pond and bioreactors up to 40.000 L (Fig. 7.5). Axenic strains, but even algal and cyanobacterial communities, are investigated; symbioses between heterotrophic/mixotrophic and autotrophic strains are desired to prolong the growth phase at low light conditions. Cyanobacteria are expected to utilize atmospheric nitrogen and convert it to ammonia, to prevent N-limitation for

Fig. 7.5: Microalgal outdoor open ponds (6000 L) located in close connection with the local combined heat and power plant (Umeå Energi), Umeå, Northern Sweden. The open ponds are inoculated mainly with locally isolated microalgal species and used for testing biomass generation, lipid productivity, and municipal wastewater treatment efficiency under local environmental conditions. (Photo: Francesco Gentili).

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algal growth in municipal and some industrial wastewaters, such as that from pulpand-paper industries, with rather low N:P ratio. The microbial biomass is then analyzed with respect to contaminants accumulating in the cells, lipids, and higher-value products. Even the use of these microorganisms as nitrogen- and phosphorous-rich feed for plants animals and fungi is investigated. In the end, a positive energy balance considering all factors from growth to harvest to refinery and end-product is required for a sustainable production; energy balances, process integration, and life cycle analyses are therefore performed. Natural wild microalgal strains with high potential for municipal wastewater remediation in Northern areas were isolated that are able to produce biomass and efficiently remove nitrogen, phosphorus, and contaminants even at low temperatures and short light periods, far superior to culture collection strains [155].

7.4.1 Why wastewater? Wastewater is a rich source of organic and inorganic compounds. It contains carbon and essential nutrients like nitrogen, phosphorus, and trace metals [156, 157]. In traditional and well-established wastewater treatment techniques, the removal of nitrogen and phosphorus is energy demanding [158]; nitrogen pollution alone costs the European Union between €70 billion and €320 billion per year [159]. On the other hand, nitrogen and phosphorus are highly important in agriculture, but the energy required to produce nitrogen and phosphorus fertilizers is high, 11.1 and 10 kWh/kg, respectively [160]. Phosphorus resources are limited and we are rapidly approaching the production peak [161]; hence, recycling this vital element is a pivotal challenge of the 21st century. Since microalgae have a high ability to take up minerals and nutrients dissolved in water, such as ammonium and phosphate, they have been used for a long time in wastewater treatments [162]. However, several challenges still need to be solved. We need algal strains able to cope with different types of wastewaters; e.g. municipal, industrial, and agricultural sewages profoundly vary in chemical and biological composition. Additionally, great variation in wastewater quality and content can be expected during the year. Photosynthetic light harvesting is also challenging; e.g., municipal and pulp and paper wastewaters are high in carbon compounds that color the water dark-brown, thus reducing the light penetration. The microalgae therefore should be adapted to low light conditions and preferentially be mixotrophic. Chlorella and Scenedesmus strains are known to grow under heterotrophic conditions and at the same time are able to remove compounds from the water with highly efficient biomass generation [163]. Research has shown that photosynthetic microorganisms are able to reduce nitrogen and phosphorus from wastewater up to 90–95% [156, 164, 165]. Green algae also have the ability to clean the urban wastewater from pharmaceutical components, like the antidepressant bupropion, the antibiotic clarithromycin, and other types of medicines [166, 167].

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The pulp and paper industry is the world’s largest producer of plant-based wastewater [168]. Even though the amount of water per ton of paper produced has been decreasing over time, between 10 and 50 m3 of water are still needed to produce a ton of paper [169, 170]. Even though the wastewater from the pulp and paper industry is rich in carbon, it is limited in nitrogen and phosphorus. Hence, in conventional wastewater treatment processes by the pulp and paper industry, nutrients had to be added to ensure microbial growth during treatment [171, 172]. Algae have been successfully used to remove chemical oxygen demands, color, and organic xenobiotics from diluted pulp and paper wastewater, with nutrients added to support the algal growth [173]. Mixing of different wastewater types has shown to improve the nutrient balance for high algal biomass production [174, 175]. Contrary to pulp and paper wastewater, sewage produced by the dairy industry is rich in nitrogen and phosphorus [176]. The dairy industry produces a volume of wastewater and sludge, which is ca 2.5 times the volume of the milk processed [177]. In agro-industrial wastewater from dairy and pig farming, algal growth efficiently removed ammonia and phosphorus [178].

7.4.2 Why flue gases? Algae grown on wastewater yield more biomass when additionally bubbled with CO2 [179]. As the atmospheric air contains only about 0.04% of CO2, increased input of CO2 to the system is necessary to have a high algal production. Bubbling algal cultures with air further on diminishes the amount of nitrogen in the media (ammonia stripping) [180]. Improving algal growth by the addition of flue gases therefore supplements the scarce inorganic carbon dissolved in the wastewater [181] and at the same time cleans the air. Flue gases are released by burning fossil-based products, i.e. coal, which even today is the most common fuel for energy production even though it greatly increases the amount of pollutants such as CO2, SOx, and NOx in the air [182]. Depending on the industry, the amount of released CO2 varies between 10% and 75%, giving rise to global temperature increase, deterioration of the ozone layer, acid rain, smog, and change of biodiversity [159] Different strategies are used to reduce the CO2 released into the atmosphere: bind it into geological formations, convert it into liquids, or produce carbonate salts [183]. So far, photosynthesis is the most efficient and the only natural process to reduce atmospheric CO2. Theoretically, to produce 1 g of biomass, microalgae and cyanobacteria have to consume about 1.88 g of CO2 [184]; experimentally, the production of 1 kg of Chlorella sp. biomass consumed up to 4.4 kg of CO2 [185]. Therefore, photosynthetic microorganisms are interesting not only from an industrial point of view but also to counteract the effects of global warming. Chlorella vulgaris was shown to be very efficient in removing both ammonia from sewage and CO2 from flue gases [186]; even other species are highly efficient in sequestering a high amount of CO2, e.g. Chlorella, Zygnema, Scenedesmus, and Chaetoceros

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species [187]; Scenedesmus obliquus and Chlorella kessleri grow at CO2 concentrations from 6% to 18% with good growth rate [188]; Chlorella even survives addition of up to 40% CO2, Scenedesmus up to 80% [189]. To be able to adapt to such high CO2 concentrations, these algae strains have to avoid acidification inside their cytoplasm and stromal compartment of the chloroplast to keep Rubisco sufficiently active [190]. Only few microalgae strains are able to survive in extreme polluted environments with high concentrations of CO2, SOx, and NOx and therefore are able to clean the flue gases also from N and S gases [191]. Monoraphidium minutum can survive in wastewater after addition of flue gases with 200 ppm of SO2 and 150 ppm of NO [192]. The addition of flue gases creates an extreme environment with very low pH, which is suitable only for very few strains of algae, but helps to avoid invasive species, competitors, and grazers. Flue gas provides not only the carbon source but also the temperature needed for optimal algal growth. Commonly, the gas must be cooled down to the optimal temperature for microalgal growth. To save this energy, algal or cyanobacterial strains able to survive in thermophilic environments (high temperature) are needed [193], like Cyanidioschyzon melorae, Cyanidium caldarim, or Galdieria partita, which showed growth at very acid environments (pH 1) and high temperatures (50oC) [194]. However, in Nordic climate, the temperature of flue gas is desired to heat the wastewater especially during autumn, winter, and spring.

7.5 Conclusions and final remarks Cold environments characterizing the Arctic, Antarctic, their corresponding subareas, and Alpine regions extensively dominate the Earth. Photosynthetic microorganisms adapted to extreme light and temperature conditions play a fundamental role being the primary producer in high-latitude ecosystems [195]. Due to their special light and temperature requirements, psychrophilic and psychrotolerant algae can thus be used as candidate models for research and commercial purposes: they are excellent examples to study the mechanisms of resource utilization and tolerance to environmental stress [195]; their cold-adapted enzymes find applications in biotechnology, in energy production, as detergent additives, in textile and food industry, and for bioremediation of cold environments [135, 196–198]. Psychrophiles further have a great potential within the field of biomass production, especially in those regions located at high latitudes, where low light and temperature pose challenges to photosynthesis. In cold climates, usually photobioreactors are placed into greenhouses and artificial light has to be used. Waste heat from a power plant may be recycled to reduce energy costs. It has been estimated that biodiesel production by algae grown in environments with low or absent natural light, low temperatures, and without a local source of waste heat consumes more energy than soy biodiesel production, and it is equally or even more polluting [199].

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The usage of extremophiles will enhance microalgal biomass production in high latitudes [14]. Sustainable local production of energy in these areas can thus be optimized by screening and characterization of algal strains for optimal growth, survival, and their potential to produce oil or other high-value molecules. In open photobioreactors, temperature can vary notably during the day and along the year: cultivation of microorganisms able to acclimate to wide ranges of temperatures represent an important advantage [34]. In a mixed culture of the two algae strains Chlorella vulgaris and Scenedesmus suspicatus, Chlorella outcompeted Scenedesmus during winter, being able to adapt and survive to cold and dark conditions and even freezing, while Scenedesmus took over during summer [200]. Mixed cultures of two or more algal species (and optionally bacteria and fungi) therefore might prolong biomass production at high latitudes, extending productivity even in critical seasons as early spring and late autumn. Photosynthetic extremophiles may even play a major role in the newly emerging field of astrobiology, where the origin, diversity, and distribution of microorganisms from the most extreme environments on Earth are studied in view of their usage for extraterrestrial life. Research on microbial communities populating polar and cold regions may not only provide interesting clues on the origin of life on Earth but also identify promising candidates for future extraterrestrial colonization [201]. A natural community of rock-dwelling phototrophs augmented with cyanobacterial cultures of Anabaenea cylindrica, Nostoc commune, and Chroococcidipsis was for example exposed to conditions of outer space for 584 days, including darkness, large temperature fluctuations, and strong UV radiation. A community consisting of the two algae Chlorella and Rosenvingiella spp., the cyanobacterium Gloeocapsa sp., and two nonphotosynthetic bacteria survived, thus confirming the great potential of extremophile microorganisms for space exploration [202].

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[132] Feller G, Narinx E, Arpigny JL, et al. Enzymes from psychrophilic organisms. FEMS Microbiol Rev. 1996;18:189–202. https://doi.org/10.1111/j.1574-6976.1996.tb00236.x. [133] Siddiqui KS, Cavicchioli R. Cold-adapted enzymes. Annu Rev Biochem. 2006;75:403–33. https://doi.org/10.1146/annurev.biochem.75.103004.142723. [134] Loppes R, Devos N, Willem S, Barthelemy P, Matagne RF. Effect of temperature on two enzymes from a Psychrophilic chloromonas (Chlorophyta). J Phycol. 1996;32:276–8. https://doi. org/10.1111/j.0022-3646.1996.00276.x. [135] Gerday C, Aittaleb M, Bentahir M, et al. Cold-adapted enzymes: from fundamentals to biotechnology. Trends Biotechnol. 2000;18:103–7. https://doi.org/10.1016/S01677799(99)01413-4. [136] Hüner NPA. Chloroplast redox imbalance governs phenotypic plasticity: the “grand design of photosynthesis” revisited. Front Plant Sci. 2012;3:255. https://doi.org/10.3389/ fpls.2012.00255. [137] Huner NPA, Öquist G, Melis A. Photostasis in plants, green algae and cyanobacteria: the role of light harvesting antenna complexes. In: Beverley R. Green, William W. Parson editors. Light-Harvesting Antennas in Photosynthesis. Springer, Dordrecht. 2003. p. 401–21. DOI: https://doi.org/10.1007/978-94-017-2087-8. [138] Ensminger I, Busch F, Huner NPA. Photostasis and cold acclimation: sensing low temperature through photosynthesis. Physiol Plant. 2006;126:28–44. https://doi.org/10.1111/j.13993054.2006.00627.x. [139] Lomas MW, Glibert PM. Interactions between NH + 4 and NO − 3 uptake and assimilation: comparison of diatoms and dinoflagellates at several growth temperatures. Mar Biol. 1999;133:541–51. https://doi.org/10.1007/s002270050494. [140] Parker MS, Armbrust EV. Synergistic effects of light, temperature, and nitrogen source on transcription of genes for carbon and nitrogen metabolism in the centric diatom Thalassiosira pseudonana (Bacillariophyceae)1. J Phycol. 2005;41:1142–53. https://doi.org/10.1111/j.15298817.2005.00139.x. [141] Fritsen CH, Priscu JC. Seasonal change in the optical properties of the permanent ice cover on Lake Bonney, Antarctica: consequences for lake productivity and phytoplankton dynamics. Limnol Oceanogr. 1999;44:447–54. https://doi.org/10.4319/lo.1999.44.2.0447. [142] Spigel RH, Priscu JC. Evolution of temperature and salt structure of Lake Bonney, a chemically stratified Antarctic lake. Hydrobiologia. 1996;321:177–90. https://doi.org/10.1007/BF00143749. [143] Spigel RH, Priscu JC. Physical limnology of the Mcmurdo Dry Valleys lakes. In: John C. Priscu editor. Ecosystem Dynamics in a Polar Desert: the Mcmurdo Dry Valleys, Antarctica. American Geophysical Union. 2013. p. 153–87. https://doi.org/10.1029/AR072p0153. [144] Howard-Williams C, Schwarz A-M, Hawes I, Priscu JC. Optical properties of the Mcmurdo Dry Valley lakes, Antarctica. In: John C. Priscu editor. Ecosystem Dynamics in a Polar Desert: the Mcmurdo Dry Valleys, Antarctica. American Geophysical Union. 2013. p. 189–203. https://doi. org/10.1029/AR072p0189. [145] Lizotte MP, Priscu JC. Natural fluorescence and quantum yields in vertically stationary phytoplankton from perennially ice-covered lakes. Limnol Oceanogr. 1994;39:1399–410. https://doi.org/10.4319/lo.1994.39.6.1399. [146] Koob DD, Leister GL. Primary productivity and associated physical, chemical, and biological characteristics of Lake Bonney: a perennially ice-covered lake in Antarctica. In: Llano GA, editor. Antarctic Terrestrial Biology. American Geophysical Union; Washington, 1972. p. 51–68. https://doi.org/10.1002/9781118664667.ch2. [147] Parker BC, Paterson RA, Craft JA, et al. Changes in dissolved organic matter, photosynthetic production, and microbial community composition in Lake Bonney, southern Victoria Land, Antarctica. In: Llano GA, editor. Adaptations within antarctic ecosystems. Gulf Pub Co, Washington, DC; 1977. p. 873–90.

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Rasheed Adeleke, Maryam Bello-Akinosho, Mphekgo Maila and Natuschka M. Lee

8 Roles of extremophiles in the bioremediation of polycyclic aromatic hydrocarbon contaminated soil environment 8.1 Introduction The presence or absence of microorganisms is critical to proper functioning of any ecosystem as they are involved in bio-geochemical cycling of many important nutrients and metals as well as biodegradation or stabilization of contaminants in the environment [1]. The discovery of microorganisms known as “extremophiles” has proven to be a major milestone in the biotechnology industry. Research studies in this field have been well received globally, especially with important discoveries that revealed life existence in extreme environments. This has led to huge investment in research studies relating to life in extreme environments by funding agencies such as the National Aeronautics and Space Administration. By definition, extremophiles are defined as organisms that thrive, grow, or survive in habitats that for other life forms are intolerably hostile or even lethal [2, 3]. Apart from studies relating to extremely cold or hot environments as well as exobiology and astrobiology, it seems that the term “extremophiles” has not been fully accepted and integrated in other aspects of biotechnology. In spite of significant amount of work devoted to microbes in contaminated environments, scientists have been cautious, or perhaps overlooked the fact that microorganisms that are capable of growing in such environments are also extremophiles. Typically, environmental conditions and edaphic parameters are factors that shape the soil microbial communities [4]. However, chronic pollution could also be an important parameter that can determine soil microbial diversity. Both the shortterm and long-term impacts of contamination could decrease microbial abundance, richness, and diversity, but the soil microbial community would adapt over time toward a unique and rich diversity pattern [5, 6]. Significant modification of natural environments can occur through the introduction of contaminants such as polycyclic aromatic hydrocarbons (PAHs) in the environment. Due to their toxic nature, PAH contamination leads to gradual and eventual favorable selection of microbial consortia that are able to adapt and survive in contaminated environments. Such consortia are able to utilize or coutilize the PAHs as carbon source [7, 8]. Such attributes of these bacteria are beneficial in environmental restoration and sustainability. This chapter describes the roles of PAH extremophiles in the biodegradation of PAHs and the impacts of PAH contamination on the soil environment. It also discusses https://doi.org/10.1515/9783110424331-008

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 8 Roles of extremophiles in the biodegradation of PAHs

the harnessing of naturally occurring plant-microbe interaction in the rhizosphere for an enhanced bioremediation of PAH polluted soil.

8.2 PAHs in soil environment The environment has been threatened by different levels of contaminants as a result of the industrial revolution witnessed in the last century. Energy generation processes associated with industrial revolution have brought about an increase in environmental PAHs as combustion of hydrocarbons from petroleum and other fossil fuels has soared high due to increased need for heat and energy generation for domestic and manufacturing purposes. PAHs are a class of widely distributed organic compounds in the environment, formed majorly as a result of incomplete pyrolysis of organic materials that contain carbon and hydrogen. They belong to the group of chemical contaminants known as persistent organic pollutants (POPs) [9]; they possess two or more fused aromatic rings that can be arranged in a linear, angular, or clustered manner [10–13]. PAHs are broadly classified into two categories depending on the number of aromatic rings, namely low molecular weight (LMW), containing three or fewer rings, and high molecular weight (HMW), containing four or more rings [14]. These molecules are highly recalcitrant and they persist in the environment due to their hydrophobicity and low aqueous solubility [15]. The chemical aromatic ring structure of PAHs confers a stabilizing delocalization or resonance energy on them and makes them inaccessible to oxidation and reduction. This further contributes to the persistence of PAHs in the environment; therefore, degradation of PAHs would only occur by an intricate degradation system [16]. The resistance of PAHs to oxidation and reduction increases with higher molecular weight, which in turn increases because of additional rings in the PAH structure. Furthermore, PAHs have been classified as “small” and “big.” When PAHs comprise up to six fused aromatic rings or less, some authors refer them as “small” PAHs, while PAHs with more than six fused rings are referred to as the “large” PAHs [17, 18]. The International Union on Pure and Applied Chemistry (http://www.iupac.org/) designates naphthalene, with only two fused rings, not a true PAH but a bicyclic aromatic hydrocarbon [17]. However, several other authors regard naphthalene a LMW PAH [14, 19–23]. The hydrophobicity of PAHs generally increases with increasing molecular mass, with aqueous solubility declining from the low mg/L range for LMW PAHs to about 1 μg/L for HMW PAHs [24]. In addition to their hydrophobic properties, they are semivolatile and usually have high affinity for sediment particles. Therefore, PAHs are readily adsorbed to soil and dust particles and could also be distributed through air [25]. Furthermore, PAHs are lipophilic and therefore soluble in organic solvents [17]. Pure PAHs are colorless to white to pale yellow-green in appearance and are solid in their purest form with a good number being fluorescent, emitting characteristic wavelengths of light when they are excited [17].

8.2 PAHs in soil environment 

 199

PAHs do not naturally occur as individual compounds but as a mixture of compounds. They are the largest class of carcinogenic compounds. Some of them are included in the European Union and US Environmental Protection Agency (USEPA) priority list of pollutants because of their mutagenicity and carcinogenicity. Sixteen PAHs are listed by USEPA as among priority pollutants (Tab. 8.1). They were so designated because more information was available about them and because there was more likelihood of exposure to them. In 1995, the Agency for Toxic Substances and Disease Registry (ATSDR) and USEPA added PAHs to their hazardous substances list, and in 2001, ATSDR ranked PAHs as the ninth most threatening chemical compounds to human health [26].

8.2.1 Direct and indirect impacts of PAH on soil environment There are considerably high levels of PAHs that can be detected in various environmental samples such as air, water, sediments, and soil [20, 28–30]. Among all the Tab. 8.1: USEPA’S 16 priority pollutant PAHs. PAH name

Number Boiling of rings point (°C)

Melting point (°C)

Molecular Uses weight (g mole−1)

Naphthalene

2

218

80.26

128.17

Acenaphthene Acenaphthylene Anthracene Phenanthrene Fluorene

3 3 3 3 3

279 265–275 340–342 340 295

93–95 92–93 218 100 116–117

154.21 152.20 178.23 178.23 166.22

Fluoranthene

4

375

110.8

202.26

Benzo[a]anthracene Chrysene Pyrene Benzo[a]pyrene Benzo[b]fluoranthene Benzo[k]fluoranthene Dibenzo[a,h] anthracene Benzo[g,h,i]perylene Indeno [1,2,3-c,d] pyrene

4 4 4 5 5 5 6

438 448 393–404 495 No data 480 No data

158 254 156 179–179.3 168.3 215.7 262

228.29 228.29 202.26 252.32 252.32 252.32 278.35

Drugs, polyvinyl chloride plastics, mothballs, toilet deodorant Dyes and pigments Research Insecticides, dyes Drug base Dyes and pigments, pesticides, thermoset plastics Dyes, pharmaceuticals, agrochemicals Research Research Research Research Research Research Research

6 6

550 530

273 163.6

276.34 276.34

Research Research

Adapted from Refs. [9, 20, 27]; ATSDR: https://www.atsdr.cdc.gov/toxprofiles/tp69-c4.pdf; http:// www.toxipedia.org/display/toxipedia/Polycyclic+Aromatic+Hydrocarbons.

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 8 Roles of extremophiles in the biodegradation of PAHs

potential destinations for PAHs, soil seems to be the ultimate sink as it contains more than 90% of all the PAHs found in the environment [21]. Soil background concentration of PAHs is often linked to atmospheric deposition, but proximity to point sources and soil properties have also been acknowledged to affect concentrations [15, 31]. Soil PAHs are also an indication of environmental PAH pollution because PAHs get enriched in soil and recalcitrantly adsorb onto soil particles [20]. Although PAHs are ubiquitous, they have strong affinity to sorb onto particulate matter such as soil and sediments. A localized PAH contamination of soil and sediment may occur in an instance of a minor spill during petroleum product transportation or refinery as PAHs constitute a substantial proportion in petroleum oil products. However, disintegration of an entire ecosystem may occur when large quantities of petroleum products are spilled into the environment. In this case, plants, animal lives, and vast quantity of microbial lives are destroyed. The enduring and surviving microbes are considered adaptable to the PAH-contaminated environment [32], being able to utilize the carbon sources in the hydrocarbons as source of energy. In the case of run-off from polluted soil to water bodies or when soil contamination is minor, there is a tendency for bioaccumulation and biomagnification of the hydrocarbons along the food chain [14, 33]. PAHs are widely accepted to be toxic to both higher animals and microorganisms. Many of the PAHs are carcinogens or synergists to other carcinogens. Some PAHs are genotoxic, mutagenic, or teratogenic. They tend to bioaccumulate in the soft tissues of higher organisms [11]. PAHs are implicated in different kinds of cancers through different exposure routes, ranging from ingestion to inhalation to dermal contact. In addition to being carcinogenic, PAHs also impair human health in various ways, with manifestations such as decreased body weight, enlarged liver with cellular edema, congestion of liver parenchyma, reproductive toxicity, intrauterine growth retardation, embryolethality, fetal malformations, learning and IQ deficits, destruction of oocytes, and inflammation of kidney cells [34].

8.2.2 Sources of PAH in soil environments Most PAHs are formed due to subsequent recombination of incompletely combusted (pyrolyzed) carbon-containing compounds at very high temperatures or not too high temperatures but for long period at suboptimum combustion conditions [17, 35]. A representation is depicted on Fig. 8.1. PAHs occur in the environment and soil largely due to human activities, but some natural phenomena contribute to their occurrence. Natural occurrences that produce PAHs are thermal geologic reactions that are associated with fossil fuel and mineral production [36], bush fires, and forest vegetation burning. Some plants and microorganisms also react in a manner that results in PAH production [12, 37, 38]. Various human activities contribute PAHs to the environment, particularly during

8.2 PAHs in soil environment 

 201

Fig. 8.1: Depiction of PAH formation from reassorted incompletely combusted C-H bonds (self-drawn).

combustion of fossil fuels, coal-tar, wood, refuse, garbage, oil filters, and used lubricating oil. PAHs could also be introduced to the soil when petroleum products are spilled in the process of transportation or dispensing [29, 35], petroleum has about 25–40% of its mass being PAHs [12, 39]. Smokes from tobacco are also a source of PAHs to the environment [40]. Broadly, the classification of PAHs based on their sources is follows [11, 39]: –– Diagenic – PAHs produced as a result of natural precursors such as volcanic eruption and microbial degradation of organic matter –– Pyrogenic – PAHs produced as a result of incomplete combustion of organic matters such as coal, oil, wood, soot, exhaust of automobiles, industrial emissions. Sometimes, forest fires, which are natural, are classified as pyrogenic. –– Petrogenic – PAHs produced from petroleum sources such as petroleum production processes and oil spill

8.2.3 Toxicity of PAHs creates extreme environment The introduction of PAHs to soil can impair the productivity of such soil by eliminating some soil organisms while some others able to utilize the carbon sources in the PAHs take advantage of the available nutrient, which may lead to the degradation of such PAHs. This may result in a shift in the microbial community and perhaps some loss of important soil functions pertaining to fertility [32]. Furthermore, the high hydrophobicity and low solubility of PAHs are limitations to their utilization and biodegradation by soil microorganisms. Therefore, only microorganisms capable of changing their outer membrane structure to facilitate direct uptake of PAHs would inhabit a PAH-contaminated soil [41]. PAH contamination thus causes soil to become extreme environment for inhabiting microorganisms; only those able to thrive would then find such polluted soil habitable. An extreme environment, impacted by xenobiotic pollutant, is thus created. Many PAH-polluted sites share similar characteristics with other extreme environments, such as extreme temperature, extreme salt concentration, and extreme pH, among others [42]. PAH extremophiles therefore tend

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to be polyextremophiles, with the ability to thrive in more than one extreme environment [2]. Several PAH polyextremophiles are expounded in the literatures, for example halophilic PAH degraders [43–46], thermophilic PAH degraders [47], psychrotolerant PAH degraders [48, 49], and acidophilic and acidotolerant PAH degraders [50, 51]. Pugazhendi et al. [52] described a halothermophilic bacterial consortium for their PAH degradation. Similarly, Hirano et al. [53] isolated and characterized a Xanthobacter species capable of PAH degradation in extremely low oxygen conditions. The toxicity of PAHs to microorganisms is sparsely reported. The degradation kinetics and the monitoring of the toxicity of environmental pollutants, such as PAHs, on microorganisms have been successfully carried out using different types of methods such as commonly used bioluminescence method [54–56]. Other more rare, but nevertheless, useful methods such as isothermal calorimetry [57] are also used in the monitoring. The robustness of the bioluminescence approach allows easy monitoring of toxicity of dangerous chemicals on microorganisms as well as direct signal measurement and the associated rapid response [55]. The luminescent bacterium that has been most extensively used in environmental toxicity is Vibrio fischeri [56]. The bioluminescence method is based on the principle that light emission intensity changes with the concentration of most toxic chemicals. The bioluminescence by microorganisms involves respiration for production of substrates needed for the generation of light. Due to the link between electron transport and the overall cell metabolism, any environmental condition that can hinder electron transport and change other related cellular processes should have an effect on bioluminescence [56, 58]. Using Escherichia coli strain harboring a lac::luxCDABE-fused plasmid, Lee et al. [54] were able to use bioluminescence for assessment and monitoring of PAH impact on microorganisms. This bacterial strain shows lower bioluminescence levels when the cellular metabolism is inhibited, hence clearly indicating that PAHs are toxic not only to humans but also to microorganisms in the ecosystem. This suggests that a PAH-contaminated environment is an extreme environment where only organisms with special attributes are able to survive and adapt.

8.3 Removal of PAHs from the environment 8.3.1 Different approaches for PAH removal from soil A number of different degradation technologies and strategies, such as physical and chemical [59], thermal [59, 60], containment [59], and biological [61] techniques, have been utilized in attempts to eliminate PAH contaminants from the environment. Physical and chemical methods of degradation have drawbacks as only being applicable to relatively small areas and unsuitable for large areas [62]. In addition, they are expensive to carry out [63]. Biological PAH degradation techniques include phytodegradation, in situ microbial degradation, landfarming, composting, and use

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of bioreactors [61, 64]. Biodegradation is defined as the elimination, attenuation, or transformation of polluting or contaminating substances by the use of biological processes [65].

8.3.2 Bioremediation of PAH-polluted sites Biodegradation, mineralization, and bioremediation are common terminologies that are widely used to describe biological remediation of polluted sites. Although these are related, they are often misused. When PAHs are degraded biologically, the complex PAH compound is broken down (metabolized) into less complex compounds, then biodegradation is achieved. If the breakdown is complete and simple compounds such as CO2 and H2O are formed, then the biodegradation is termed mineralization [66]. Bioremediation is a term used when organisms utilized in biodegradation and mineralization are used in the elimination or attenuation or transformation of pollution in an environment. Biodegradation is therefore a mechanism in accomplishing bioremediation [67]. Numerous PAH-degrading microbial strains have been isolated and characterized in recent years. This development matches the improving level of technology that has yielded information about the abilities of such microorganisms and their consortium to biodegrade many more PAHs, as well as elucidation of the biodegradation pathways. In spite of this advancement, there are still a significant number of challenges that are yet to be unraveled that relate to the regulatory mechanisms of PAH biodegradation, simultaneous biodegradation of PAHs with other hydrocarbons in mixtures, as well as the microbial interactions within PAH-degrading consortia [68]. Phytoremediation is a prominent technique employed in bioremediation of PAHs, an in situ technique that utilizes living green plants for removal, degradation, or containment of contaminants in different sites [69, 70]. It involves a number of processes such as: –– Phytodegradation, which is the use of plants to uptake, store, and or degrade contaminants within the plants tissues. –– Phytostimulation or rhizodegradation, which is the use of rhizospheric associations between plants and soil microorganisms to degrade contaminants. –– Phytovolatilization, which is the use of a plant’s ability to uptake contaminants from the growth matrix, then transform, and volatilize such contaminants into the atmosphere. –– Phytoextraction, which involves the accumulation of toxic compounds in the harvestable part of a plant. The plant parts are incinerated after harvesting and the ashes are either treated as hazardous residues or phytomined for the recovery of the metals. –– Phytostabilization, where the plant itself or the root exudates help to sequester the soil contaminant, thus making it less available.

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Plants are beneficial in bioremediation because plant root deposits provide energy, which drives microbial degradation process, thus allowing for diauxic growth. Roots are also able to alter the physical and chemical conditions in polluted soil such that microbial degradation is promoted [71]. When plants and their root-associated rhizospheric bacteria are involved in the clean-up of polluted terrestrial environments, the plants assist in stabilizing bacterial performance by providing a specific ecological niche and nutritionally favorable conditions to ensure maintenance of bacterial cell numbers and activities [72]. A variety of grasses and trees have been used in phytoremediation, including Zea mays (maize), Brassica napus (canola), Elsholtzia splendens, Oenothera biennis, cannabi, mustard, Hibiscus, Amaranthus, Festuca arundinacea, and a variety of pine including Aleppo pine (Pinus halepensis Mill) [73]. As recalcitrant as PAHs are, a variety of bacteria, fungi, and algae have been isolated and characterized for their ability to utilize and degrade PAHs. A number of studies report uncultured bacteria as also capable of PAH degradation. For instance, Eriksson et al. [74] reported an unculturable community of bacteria capable of biofilm production for the degradation of pyrene. Unculturable (better put, yetto-be-­cultured) soil bacteria far outnumber the culturable ones; studies have suggested that of all soil bacterial population, only 1–10% have so far been cultured and described [75, 76]. This may suggest that yet-to-be-cultured PAH degraders could be significantly important in the remediation of polluted soils. Tab. 8.2 gives a summary of the different taxa of 2674 culturable and 9836 unculturable archae, bacteria, and fungi retrieved from the database of SILVA (version 128, https://www.arb-silva.de) for the degradation of PAHs. Tab. 8.2: Taxa of cultured and uncultured PAH degraders (survey based on data in the Silva database, version 128, https://www.arb-silva.de). Taxa Asconditabacteria (SR1) AC1 Acidobacteria Actinobacteria Aminicenantes Archaea Armatimonadetes Atribacteria Bacteriodetes BRC1 Caldiserica Candidatus Chlamydiae Chlorobi Chloroflexi Cloacimonetes Cyanobacteria

Number 1 2 385 1355 19 182 10 1 583 3 1 6 1 13 259 2 139

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Tab. 8.2 (continued) Taxa Deferribacteres Deinococcus Dependentiae (TM6) Elusimicrobia Eukaryota FCPU426 Fibrobacteres Firmicutes Fusobacteria Gemmatimonadetes GN01 Gracilibacteria Hydrogenedentes Ignavibacteria Latescibacteria Lentisphaerae Marinimicrobia Microgenomates Nitrospinae Nitrospira Omnitrophica Parcubacteria Peregrinibacteria Planctomycetes Proteobacteria Sacchribacteria SBR1093 Spirochaetae Synergistics Tectomicrobia Tenericutes Thermatotrogae Unidentified Verrucomicrobia WS2 Zixibacteria (RBG-1)

Number 15 3 9 6 148 1 8 550 33 91 1 4 2 61 17 4 1 15 15 98 14 17 7 194 8054 14 2 35 1 1 17 2 2 102 1 3

8.4 How do extremophiles drive the bioremediation process? Extremophiles generally degrade PAHs aerobically or anaerobically. Extremophiles in PAH-contaminated environment are suggested to have co-evolved with naturally occurring compounds that have comparable chemistry to PAHs. In addition, exposure to PAH contamination for long periods may also confer such capabilities and adaptation on PAH extremophiles [77]. PAH extremophiles can be prokaryotes or

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eukaryotes; they possess special capabilities that enable them to respond to environmental signals such as sudden increase in PAH concentration. This type of specific stress response is associated with their abilities to temporarily change their mode of gene expression. Studies have reiterated and emphasized the importance of bacteria as the most important extremophiles in bioremediation [13, 23]. A variety of genera of Gram-positive and Gram-negative bacteria have been described for their ability to utilize and degrade PAHs; a summary is given in Tab. 8.3. In the fungal kingdom, the white-rot fungi [78, 79] and some species of Penicillium, Trichoderma, and Aspergillus [80] have been found to degrade different PAHs. Ectomycorrhizal fungi (ECMF) alone or in association with ectomycorrhizal plants have also been found to degrade PAHs [81]. Fungal exoenzymes are suggested to initiate the degradation of HMW PAHs, diffusing toward PAHs and interacting with them, thereby facilitating PAH degradation by bacterial communities [18, 82]. Few bacterial cells can survive and grow in the presence of HMW PAHs. This is probably due to the high retention of these compounds on the solid soil phase, resulting in mass transfer rates of HMW PAHs to the bacterial cells, which are often too low for the basic cell metabolite requirements. In addition, this could also be the potential reason for lack of evolution of suitable enzymatic pathways for biodegradation of HMW PAHs [83]. Another important attribute of PAH extremophiles is their ability to excrete biosurfactants, bioemulsifiers, coenzymes, extracellular polymeric substances, or their formation of biofilms. With any of these, PAH extremophiles can increase the solubility and bioavailability of PAHs [82, 84–86]. It is also important to mention that PAH extremophiles are able to show chemotaxis toward PAHs. This is an important step in biodegradation as it reduces the distance between the PAH extremophiles and the contaminant [8, 18, 85, 87]. Bioremediation of PAH-polluted soils is influenced by factors such as those pertaining to the soil – soil type, pH, temperature, aeration, moisture content, LMW organic acids and humic acids, redox potential, and inorganic nutrient availability [21, 88–90]. A predominant factor in bioremediation is the concentration and inherent properties of the polluting PAH, while most important is the organisms employed in the remediation process [89, 90]. During the course of soil PAH biodegradation, a range of metabolites is produced, some of which are environmentally friendly while some others are not. Non-environmentally friendly biodegradation products are sometimes result of biodegradation by some fungi and are themselves an environmental problem. Research results have shown that PAHs are mineralized in cocultures of bacteria and fungi where the intermediate metabolic product from the fungi serves as the substrate for bacterial mineralization [22]. In spite of claims from many studies about the abilities of both eukaryotes and prokaryotes in soil biodegradation of PAHs, the rate of removal is usually very slow. This means that it is important to optimize and design improved methods of bioremediation of PAHs in the environment using the available knowledge and information. A process

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Tab. 8.3: Examples of isolated archaea, bacteria, and fungi associated with PAH degradation. Microorganism

Domain

Reference

Haloferax spp. Halobacterium sp. Halococcus sp. Haloarcula sp. Acidobacterium spp. Thermotomaculum spp Candidatus solibacter Micrococcus luteus Kocuria rosea Illumatobacter Rhodococcus sp., Corynebacterium spp Mycobacterium spp. Candidatus Microthrix Gordonia spp. Brevibacterium sp. Actinotalea sp. Cullulomonas sp. Microbacterium spp Salinibacteria spp. Athrobacter spp. Kocuria spp. Micrococcus spp. Sinomonas sp. Amycolatopsis sp. Salegentibacter sp. Flavobacterium sp. Muricauda spp. Bacillus spp. Paenibacilus spp. Staphylococcus spp. Pseudomonas spp.

Archaea Archaea Archaea Archaea Acidobacteria Anaerobic Acidobacteria Acidobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria Actinobacteria Bacteriodetes Bacteriodetes Bacteriodetes Firmicutes Firmicutes Firmicutes Proteobacteria

Ochrobactrum spp. Hyphomonas sp. Brevundimonas sp. Methylobacterium spp. Rhizobium sp. Parvibaculum sp. Labrys sp. Xanthobacter sp. Celeribacter spp. Labrenzia aggerata Paracoccus spp

Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria

[92–95] [95] [95] [94] [96, 97] [98] [99] [100] [100] [98] [101–104] [105] [105–114] [105] [105, 115, 116] [117] [105] [105] [118–120] [117] [121, 122] [123, 124] [120, 125–128] [129] [130] [120] [131–133] [120, 134] [117, 120, 122, 127, 132, 135–140] [132, 141] [96, 127, 132, 134, 142] [96, 104, 117, 118, 120, 123, 127, 131, 132, 143–159] [126, 131, 139, 152, 160–162] [163] [164] [122, 165] [119, 122, 126, 132, 166] [134, 163] [104, 131] [53] [101, 167–169] [120, 163] [134, 170]

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Tab. 8.3 (continued) Microorganism

Domain

Reference

Thalassospira sp. Tistrella spp. Erythrobacter sp. Novosphingobium Sphingomonas sp. Sphingopyxis sp. Achromobacter spp. Alcaligenes spp Pusillimonas spp. Burkholderia sp Ralstonia sp. Acidovorax sp. Camamonas sp. Delftia sp. Rubrivivax sp. Varivorax Oxolobacter sp. Masillia sp Aeromonas sp. Alteromonas sp. Marinobacter sp Idiomarina sp. Shewanella sp. Porticoccus sp. Enterobacter sp. Klebsiella sp. Pantoea sp. Alcanivorax sp. Halomonas spp. Marinobacterium sp. Acinetobacter spp Cycloclasticus sp. Methylophaga sp. Algiphilus sp. Dyella sp. Luteibacter Pseudoxanthomonas spp. Stenotrophomonas sp. Novosphingobium sp. Aspergillus spp. Fusarium spp. Trichoderma sp. Saccharomyces spp.

Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Proteobacteria Fungi Fungi Fungi Fungi

[46, 120, 163, 171, 172] [120, 170, 173] [120, 162, 173, 174] [118, 125, 162, 173, 175–177] [120, 131, 133, 175, 178–182] [175] [126, 132, 163, 183, 184] [118, 120, 131, 162, 185] [118] [112, 133, 162, 186] [104, 122, 133] [187] [112, 133] [112] [188] [153] [112] [189, 190] [132] [191, 192] [120, 134, 173] [46, 120, 134] [118, 193, 194] [162, 195] [196] [197] [127, 132] [43, 46, 120, 125, 163, 173] [43, 44, 46, 120, 163, 173] [162, 173, 198] [121, 122, 127, 134, 193, 199, 200] [151, 158, 201, 202] [134] [203] [162, 204] [162] [121, 131, 205, 206] [112, 121, 126, 131, 207, 208] [209] [100] [131, 210, 211] [212] [213]

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 209

enhancer that could be exploited is aeration of soil to facilitate improved aerobic degradation of pollutants. This could be achieved mechanically or by the introduction of pollutant-tolerant earthworms [91].

8.4.1 Aerobic and anaerobic degradation processes When PAHs are degraded aerobically, microorganisms either degrade them as a sole source of carbon and energy (metabolism) or by cometabolism. Cometabolism is a process where an enzyme produced by the microorganism for degradation of certain PAH also degrades another PAH that is not used or not essential for its growth [24]. This process is especially important for the degradation of mixtures of PAHs. The initial step in aerobic degradation is the incorporation of oxygen into the PAH structure by enzymes called oxygenases produced by microorganisms, which catalyze oxygen-fixing reactions. This is the rate-limiting step in the biodegradation of PAHs [7]. Oxygenases may be monooxygenases, generally produced by fungi, which incorporate only one oxygen atom into the substrate to form arene oxides, or they may be dioxygenases, characteristically produced by bacteria. Dioxygenases incorporate two oxygen atoms into the substrate. The incorporation of the two oxygen atoms to two adjacent carbon atoms of the aromatic ring is referred to as a dihydroxylation process and it is catalyzed by a ring-hydroxylating dioxygenase (RHD) [214]. RHDs are a large family of enzymes that are diverse with regard to their substrate specificity and protein sequence [215]. Dihydroxylation of adjacent carbon atoms of the aromatic ring lead to the formation of dioxethanes that are further oxidized to cis-dihydrodiols and then to dihydroxyl products [24]. A cis-hydrodiol is further dehydrated by a dehydrogenase enzyme, then a dihydroxylated intermediate called catechol, is formed. Degradation of catechol ultimately yields tricarboxylic acid (TCA) cycle ­intermediates. These steps are depicted in Fig. 8.2. RHDs are bacterial metalloenzymes that catalyze the first step in the biodegradation of a variety of aromatic hydrocarbons [216]. They are a multicomponent enzyme consisting of an iron-sulfur flavoprotein reductase (or ferredoxin reductase or oxidoreductase), an iron-sulfur ferredoxin, and an iron-sulfur protein (ISP), which is a dioxygenase that bears the active site of the enzyme that interacts with aromatic compounds. The ISP is composed of large alpha (α) subunits and small beta (β) subunits. The dioxygenase is most often a heterohexamer having a α3β3 configuration. A structural study of representatives of dioxygenases shows that the α-subunit is composed of two conserved domains, which are a Rieske domain and a catalytic domain. The Rieske domain binds a [2Fe-2S] cluster and the catalytic domain that contains the substrate-binding site and a mononuclear non-heme Fe center [217], where oxygenation of the substrate takes place [218]. The catalytic domain determines substrate specificity.

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Fig. 8.2: Representation of the steps involved in aromatic ring degradation (self-drawn).

Although slower and less practiced, anaerobic degradation of PAHs takes place in anoxic environmental circumstances, performed by anaerobic microorganisms. Anaerobic degradation of PAHs has been demonstrated in studies under methanogenic conditions as well as where nitrate, ferric iron and sulfate serve as electron acceptors [148, 219–221]. Anaerobic degradation of LMW PAHs is more often reported, with fewer reports of anaerobic degradation of HMW PAH [222–224]. In addition, Liang et al. [221] recently demonstrated the degradation of HMW benzo(a)pyrene and fluoranthene with nitrate serving as the electron acceptor, using a facultative ­anaerobic Pseudomonas strain. Similarly, Yan et al. [225] also investigated anaerobic degradation of benzo(a)pyrene and pyrene while organic matter derived from

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cyanobacteria was used to amend the polluted sediment. Anaerobic biodegradation is common in contaminated sediments and groundwater that tend to become anaerobic due to high pollutant concentration and associated biodegradation processes [221]. Submerged soil, for example rice paddy soils, saturated with water while air is excluded would also become anoxic and therefore require anaerobic degradation of inherent pollutants [219].

8.4.2 Bioremediation technologies Bioremediation technologies are broadly categorized into two, and these involve ex situ and in situ technologies. Ex situ technologies are usually on large scale and may involve construction of biopiles or windrows, which can be on site or a remote location [226, 227]. Usually involved is the physical removal of the contaminated material from the point of contamination [228]. In situ technologies involve soil treatment in place and are usually with fewer earthworks and may not be as effective as ex situ technologies due to lack of control and a longer decontamination duration [226]. Aerobic procedures are usually applied in situ soil bioremediation, more often via bioventing. Some examples of in situ bioremediation technologies are phytoremediation, bioventing, pump and treat, and intrinsic bioremediation. Ex situ bioremediation technologies are faster and easier to control. Examples are landfarming, bioreactors, bioventing, biofilters, and composting [228]. Important challenges associated with bioremediation technologies relate to the slowness of the method as well as the accelerating number of polluted environments. These challenges are complicated by societal complacency as people sometimes think soil is an infinite resource. As recommended by Gillespie and Philp [226], it is necessary to preserve soil for future generations in the context of bioeconomy and sustainability. Potential solutions to these challenges, especially the typical slow rate of bioremediation, are biostimulation and bioaugmentation approaches. Biostimulation involves the use of nutrient enhancement to improve the growth of indigenous PAH extremophiles [228]. For bioaugmentation, there is utilization of the genetically modified PAH extremophiles into the contaminated environment [18]. In spite of the adoption of these two approaches, extreme caution and professionalism are needed to implement the methods as they both have limitations. For instance, there are potential risks associated with bioaugmentation with genetically modified PAH extremophiles. This is similar to risks with other genetically modified organisms, and caution should always be exercised in introducing them into the environment [18, 226]. Microbial bioremediation can be a more successful technique if it would reveal beyond just the identification of microbes and catabolic genes present in contaminated soil sample. In addition, the technique should aptly elucidate and predict the threshold of pollutants that are detectable by microbes for the induction of expression of their catabolic pathways [226].

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8.5 Future perspectives 8.5.1 Can other plant-associated microorganisms – endophytes, plant growth promoting rhizobacteria (PGPR), mycorrhizal helper bacteria, and mycorrhizal fungi – assist extremophiles in PAH degradation? It is certain that the ability of extremophiles to adapt, survive, and utilize PAHs in PAH-contaminated environments may not be enough to drive the bioremediation process at the required speed and efficiency. There are a growing focus and realization that for bioremediation to be efficient, a combination of organisms is needed. This has spurred interests on the potential use of plants to remediate soils contaminated with PAHs [229–231], termed phytoremediation. Phytoremediation is a term that is often used to describe the use of plants to remediate both heavy metal and POP-polluted soils. However, phytoremediation has been used interchangeably in some literature with rhizosphere remediation (rhizoremediation). Based on the suggestion of Meharg and Cairney [232] regarding these two terms, rhizosphere remediation is a more appropriate term when plants are used to remediate POPcontaminated sites because this process occurs in the rhizosphere rather than in the plant. Remediation taking place in the rhizosphere is advantageous as it prevents the transfer of the pollutants from soil to the plants and the wild-life food chains [229, 232, 233]. Studies have shown that plant roots host different microbes within their rhizosphere and endosphere. Some of such microbes possess important catabolic genes for hydrocarbon degradation [234, 235]. Although remediation processes can be attributed to many different activities occurring in the rhizospere, plant-stimulated microbial activity is one of the most important. Other biological and physical factors such as bacterial plasmid transfer, transpiration stream, and alteration of soil structure are also very important to successful remediation of contaminated rhizosphere soils. Rhizosphere-associated microorganisms may not degrade POPs to yield energy, but they may cometabolize them as a result of using plant-derived cyclic compounds for their energy source. For instance, catechin and coumarin from plants serve as cometabolites during the bacterial degradation of polychlorinated biphenyls [229]. A similar scenario was illustrated by Sandmann and Loos [236], who suggested that some rhizosphere microorganisms can utilize the same enzymes for degradation of plant-derived compounds and POPs. They arrived at this conclusion because of their isolation of 2, 4, dichlorophenol (2,4-D)-degrading bacteria from the rhizospheres of sugarcane soil that was 2,4-D-free. It is important to pay more attention to the issue of cometabolism among the rhizosphere microorganisms as it is evident that the enzymatic processes needed to degrade some PAHs may not be possessed by a single organism [232]. Rhizosphere remediation of PAH-contaminated sites comes with different challenges. Usually, contaminated sites have more than one contaminant; therefore, selected plant species must be resistant to all contaminants.

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Importance and limitation of bacterial extremophiles in rhizosphere remediation of PAHs Although there are significant numbers of bacteria that have been discovered with potentials to mineralize or degrade PAHs, there are still some critical factors that affect the exclusive use of bacteria in rhizosphere remediation [237]. Bacteria are sometimes unable to exhibit enzymatic activities for complete mineralization of PAHs, especially the HMW PAHs. As reported by Fernández-Luqueño and coworkers [18], a possible reason may be the high retention of these compounds by the solid soil phase resulting in mass-transfer rates of HMW-PAHs to the bacterial cells, which are too low to match the cells’ basic metabolic requirements. Such a low ­bioavailability of PAHs may have prevented the evolution of suitable enzymatic pathways in soil bacteria. Furthermore, another limitation of the use of only bacteria extremophiles for rhizosphere remediation is the uneven distribution of the bacterial cells in the contaminated area of the soil [72, 238, 239]. Thus, when bacteria are used, there is a need for constant turning and mixing of the soil to ensure proper distribution of the bacterial cells within the contaminated zone. This process is not always easy and automatically adds to the cost of the remediation. A viable option to deal with the above-mentioned challenges is to consider cometabolism. Through cometabolism, microorganisms can degrade PAHs (usually more complex and recalcitrant) other than the one from which they derive their carbon and energy source [23]. The cometabolism process has been described as a nonspecific enzymatic reaction with a substrate that is competing with a structurally similar primary substrate for the enzyme’s active site. For example, the cometabolism of benzo[a]pyrene by a strain of Stenotrophomonas maltophilia growing on pyrene was reported by Boonchan et al. [22]. The strain had previously mineralized pyrene as a sole carbon and energy source. Likewise, in the presence of phenanthrene, Sphingobium sp. strain PNB could cometabolize acenaphthalene, benz[a]anthracene, fluoranthene, pyrene, and benzo[a]pyrene, which individually could not serve as its growth substrate [240]. Importance and limitation of ECMF in Rhizophere remediation of PAHs Fungi have been found to be potentially useful in the bioremediation processes. Perhaps, the most investigated fungi with bioremediation potentials are the white rot fungi (WRF) and mycorrhizal fungi, which could be arbuscular mycorrhizal or ECMF. There are considerable similarities between these two groups of fungi; both WRF and ECMF produce extracellular oxidative lignin and humic acid degrading enzymes [241]. Although both WRF and ECMF can play similar roles in forest ecosystems, there is a sharp contrast in their ecological functions. ECMF are symbionts of tree roots, while WRF are saprotrophs, degrading dead plant derived materials. With more than 6000 species of ECMF worldwide, only few have been investigated for their bioremediation potential. There is supporting evidence about the

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ability of ECMF to produce a range of enzymatic activities that allow, at least, partial metabolism of soil organic compounds and POPs [242, 243]. Important biodegradative enzymes that include tyrosinase, catechol oxidase, peroxidases, ascorbate oxidase, and laccase have been discovered to be associated with symbiotic ECMF mycelia in unsterile soil [244, 245]. Emerging facts from research studies appear to have concluded that ECMF are unable to completely mineralize PAHs, but there seems to be an undisputed acceptance of the fact that ECMF can metabolize and degrade PAHs [71, 72, 232, 246]. Rhizosphere bacteria can be helpful in establishing a stable relationship in the mycorrhizosphere, where they may cometabolize pollutants while utilizing the plant-derived compounds [232]. Consistently, positively influencing the growth of ECMF are a group of bacteria, referred to as mycorrhization helper bacteria [247]. They colonize the mycorrhizae, improve mycorrhizal formation with plants, and thereby improve both plant and ECMF growth. This relationship could imply an improvement to the PAH-degradation ability of ECMF partaking in remediation. Combination of fungi and bacteria in rhizosphere remediation of PAHs contaminated soil – ECMF and bacteria The use of a concerted approach that will involve plants, as well as fungal and bacterial extremophiles, may represent a solution to the challenge of incomplete mineralization of PAHs in our environment. The benefit of using fungi such as ECMF in rhizosphere remediation is directly linked to their ability to obtain their carbon supply from their plant hosts to support growth into contaminated or polluted substrates. Some of this carbon may subsequently be available to bacteria associated with the mycorrhizal mycelium [248], and this can be very significant for remediation in the mycorrhizosphere. The use of the combined approach (plant-mycorrhizal fungi-bacteria) will therefore create a scenario where both bacteria and the ECMF can play mutual roles in cometabolizing PAH contaminants. The external mycelial phase of ectomycorrhizae can spread rapidly after transplantation of mycorrhizal plants [249]. This can directly improve the efficient distribution of mycorrhizosphere-­ associated bacteria without the need for excavation or soil mixing. In addition, such practice may effectively eliminate the traditional method of soil nutrient amendment needed for maintaining active microorganisms during rhizosphere remediation [72]. Furthermore, the bacterial extremophiles could also benefit indirectly from the plant-associated microorganisms known as endophytic bacteria, or simply endophytes. Endophytes are nonpathogenic bacteria that inhabit internal plant tissues, without causing symptoms of disease [250, 251]. The major entry route of endophytes into the plant system are the roots, from where they can either localize or be disseminated to other parts of the plants such as shoots, stems, and leaves [252]. Therefore, there is usually a higher population of endophytes in the roots than other parts of the plant [250]. Exploring the interaction between plants and endophytic bacteria has

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been suggested to enhance the efficiency of remediation of pollution in soil [253, 254]. In the plant-endophyte bacteria interaction, the plant provides habitation and nutrient source to the bacteria. In return, the bacteria may act to improve plant growth and enhance stress tolerance [255], thereby indirectly aiding the detoxification process in the rhizhosphere through their influence on plants and extremophiles. Endophytic bacteria also interact more intimately with their host as they naturally colonize the host tissues; thus, their interaction with the host plant is suggested to be more than that of rhizobacteria. There are recent reports of isolated endophytes with pollutant-degradation and plant growth-enhancing abilities from different plants [235, 256]. For instance, Khan et al. [257] described degradation of phenanthrene by the duo of Willow plants and Poplar grasses in combination with a Pseudomonas species. The endophytic Pseudomonas species was found to confer resilience and stress resistance on the plants and grasses as opposed to those plants and grasses that were not inoculated with endophytes, which wilted off with chlorotic leaves. Similar reports of pollutant degradation and enhanced plant survival in the face of pollution were also reported by Phillips et al. [258], where different plant species including Alfalfa, Rye grass, Altai wild rye, and Nuttal’s salt meadow grass were used and the predominant endophytes were Pseudomonas and Brevundimonas species. A similar scenario of reduction of phenanthrene accumulation in ryegrass planted on phenanthrene contaminated soil was reported by Sun et al. [259]. The ryegrass had earlier being inoculated with a strain of phenanthrene-degrading endophytic Pseudomonas species. Likewise, a consortium of eight PAH-degrading endophytes were also found to reduce accumulation of PAHs by vegetables when grown on contaminated sites [260]. These two later reports have interesting positive health implications. In addition, pollutant-­ degradation gene expression by endophytes suggests their important contribution to organic contaminant mineralization in the soil. Endophytes have been suggested to be important for in planta hydrocarbon degradation as they may capacitate plants with resources necessary for detoxification [235]; thereby, such plants will survive adequately in contaminated environment [256]. Endophytes are also a well-known group of plant growth promoting bacteria, which can stimulate plant growth, increase yield, reduce pathogen infection, and alleviate plant stress [261, 262]. In addition, endophytes are generally known to participate in the metabolic pathways of their host as a result of long-term association with their host [263]. As a result, endophytes may benefit some genetic information from their host, which may facilitate the production of specific biologically active compounds [264, 265]. The exchange of genetic information was suggested by Afzal et al. [255] to be through horizontal gene transfer, which has long been hypothesized. In difference to this mechanism of transfer, Taghavi et al. [251] earlier demonstrated that the horizontal gene transfer is rather between and among microbial endophytes, rather than between host plants and endophytes. Endophytes have been shown to be able to produce the same or similar secondary metabolites as their host, which

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can then be transferred horizontally between them. Mechanisms were proposed for the simultaneous production of these biological compounds, and in some cases, the biosynthetic mechanism of the same compound evolves independently in plants and their microbial counterparts [266]. It has been strongly suggested, however, that interactions between endophytes and their respective plant host contributes to the co-production of these bioactive molecules [267], which could be of great importance to bioremediation of PAHs.

8.6 Summary and Conclusions Soil polluted with PAHs presents an extreme environment due to the very high toxicity of some PAHs. This is a case of a chemically extreme environment, which is often overlooked as an extreme environment. The PAH extremophiles are environmentally useful; they are able to enzymatically break up the bonds in the pollutant and utilize the carbon as energy source for their survival. The PAH degraders thrive where others organisms are unable to; they could be bacterial, fungal, or archaeal. They could act alone or as a consortium or in mutualistic association with other organisms. Oftentimes, they are polyextremophiles, with ability to thrive in more than one extreme environment. Soil bioremediation by PAH extremophiles is usually a slow one. The way forward is exploiting biostimulation, bioaugmentation, and the natural relationship of the degraders with other organisms such as the rhizospheric organisms, endophytes, phyllospere bacteria, and their plant hosts to enhance the biodegradation process. Endophytes intimately interact with host plant and participate in the metabolic pathways of their host. This interaction could be well exploited in rhizosphere remediation to enhance PAH extremophiles in remediation. Phyllospere bacteria are those bacteria found on the aerial parts of plants; much less research has been conducted about them in past decades. This is an area of research that has just begun to receive attention. It will be interesting to find out how phyllosphere bacteria would also play an assistive role in the remediation of polluted soil. The role of biodegraders in polluted soil is a very important one; it is one leading to a cleaner and greener environment in a cost-effective way.

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[253] Khan S, Afzal M, Iqbal S, Khan QM. Plant-bacteria partnerships for the remediation of hydrocarbon contaminated soils. Chemosphere. 2013;90(4):1317–32. [254] Yousaf S, Afzal M, Reichenauer TG, Brady CL, Sessitsch A. Hydrocarbon degradation, plant colonization and gene expression of alkane degradation genes by endophytic Enterobacter ludwigii strains. Environ Pollut. 2011;159(10):2675–83. [255] Afzal M, Khan QM, Sessitsch A. Endophytic bacteria: prospects and applications for the phytoremediation of organic pollutants. Chemosphere. 2014;117:232–42. [256] Oliveira V, Gomes N, Almeida A, et al. Hydrocarbon contamination and plant species determine the phylogenetic and functional diversity of endophytic degrading bacteria. Mol Ecol. 2014;23(6):1392–404. [257] Khan Z, Roman D, Kintz T, delas Alas M, Yap R, Doty S. Degradation, phytoprotection and phytoremediation of phenanthrene by endophyte Pseudomonas putida, PD1. Environ Sci Technol. 2014;48(20):12221–8. [258] Phillips LA, Germida JJ, Farrell RE, Greer CW. Hydrocarbon degradation potential and activity of endophytic bacteria associated with prairie plants. Soil Biol Biochem. 2008;40(12):3054–64. [259] Sun K, Liu J, Gao Y, Sheng Y, Kang F, Waigi MG. Inoculating plants with the endophytic bacterium Pseudomonas sp. Ph6-gfp to reduce phenanthrene contamination. Environ Sci Pollut Res. 2015;22(24):19529–37. [260] Wang J, Liu J, Ling W, Huang Q, Gao Y. Composite of PAH-degrading endophytic bacteria reduces contamination and health risks caused by PAHs in vegetables. Sci Total Environ. 2017;598:471–8. [261] Compant S, Clément C, Sessitsch A. Plant growth-promoting bacteria in the rhizo-and endosphere of plants: their role, colonization, mechanisms involved and prospects for utilization. Soil Biol Biochem. 2010;42(5):669–78. [262] Mitter B, Brader G, Afzal M, et al. Advances in elucidating beneficial interactions between plants, soil and bacteria. Adv Agron. 2013;121:381–445. [263] Golinska P, Wypij M, Agarkar G, Rathod D, Dahm H, Rai M. Endophytic actinobacteria of medicinal plants: diversity and bioactivity. Antonie van Leeuwenhoek. 2015;108(2):267–89. [264] Chithra S, Jasim B, Sachidanandan P, Jyothis M, Radhakrishnan E. Piperine production by endophytic fungus Colletotrichum gloeosporioides isolated from Piper nigrum. Phytomedicine. 2014;21(4):534–40. [265] Rai M, Agarkar G, Rathod D. Multiple applications of endophytic Colletotrichum species occurring in medicinal plants. Nov Plant Bioresour Appl Food Med Cosmet. 2014:227–36. [266] Bömke C, Tudzynski B. Diversity, regulation, and evolution of the gibberellin biosynthetic pathway in fungi compared to plants and bacteria. Phytochemistry. 2009;70(15):1876–93. [267] Heinig U, Scholz S, Jennewein S. Getting to the bottom of Taxol biosynthesis by fungi. Fungal Divers. 2013;60(1):161–70.

Guðný Vala Þorsteinsdóttir and Oddur Vilhelmsson

9 Bioremediative potential of bacteria in cold desert environments 9.1 Bioremediation – general considerations With the opening up of the Arctic to shipping, petroleum extraction, tourism, and other human activities, the risk of pollutant release into Arctic environments is steadily increasing [1]. This has, in recent years, led to increased awareness among the northern countries of the need for environment-friendly processes for pollution control and cleanup, which are now being sought after to replace less eco-friendly practices in many industries, as well as on the municipal level [2]. Ironically, the act of cleaning up pollutants from the environment can in some cases be harmful to the environment due to physical and/or chemical disturbance. Bioremediation, that is, the degradation, neutralization, or removal of contamination by living organisms, is potentially a less environmentally harmful way of cleaning up environmental pollution. It should be stressed, however, that bioremediation is, even under the best of circumstances, a very slow process compared with physical methods and is thus almost always used in conjunction with physical pollution removal in real-world situations [1]. There are multiple technical strategies with which one can approach bioremediation, which can be broadly sorted into in situ techniques and ex situ techniques. Bioremediation in situ is when the pollution is attacked at the polluted site in its “natural environment,” whereas ex situ bioremediation is handled offsite at purpose-built facilities such as biopiles or bioreactors. For in situ bioremediation, the two main strategies thought to be the most promising ways to bioremediate soil and groundwater are bioaugmentation and biostimulation techniques [3]. Biostimulation refers to the techniques employed when the indigenous microbial community is stimulated to overcome barriers in metabolic pathways and limitations of degradation. That can be attained by providing nutrients or other compounds that enhance the biodegradation in any way, such as oxygen or other electron receptors. Bioaugmentation is the process of introducing allochthonous microorganisms into the environment, usually because they are better degraders than those of the autochthonous microbiota. These technical considerations for bioremediation in general have been reviewed multiple times [3–8], and we refer the reader to these reviews for an indepth discussion. Bioremediation fundamentally rests on the process of biodegradation, whereby an organism depolymerizes and oxidizes an environmental organic compound or macromolecule with the terminal products being carbon dioxide and water, a very

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common process and a major component of the global carbon cycle [9]. This process of microbially mediated decomposition of substances can result either from the microorganism’s utilization of a certain compound as a carbon and energy source or of the organism’s co-metabolism of that compound [10], leading to a staggering versatility of biodegradation activities within the microbial world. Indeed, the maxim often attributed to C.B. van Niel [11], that for any naturally occurring compound there exists an organism capable of degrading it, can, broadly speaking, be taken as true. Or, to put it another way: “given favourable environmental conditions, all natural organic compounds degrade in the end” [12] and biodegradation by naturally occurring microbial populations is indeed important ecologically. Nevertheless, bioremediation efforts are often confounded by a disparity in the biodegradation behavior of organisms in natural environments versus in laboratory culture, or between natural environments of differing characteristics [13]. When initiating bioremediation of a certain pollutant, either in the laboratory or in the natural environment, the degradation of that pollutant needs to be favorable; that is, the intermediates and products of the degradation pathway must be at least less toxic than the substrate, preferably neutralized or completely degraded.

9.2 Hydrocarbon degradation Hydrocarbons are the most widespread pollutants and are of great concern worldwide because of their toxicity and persistence in the environment [5]. Due to their chemical inertness and the high energy barrier that needs to be overcome in order to cleave the apolar C-H bond, microbial degradation of hydrocarbons is intrinsically challenging. Nevertheless, degradation of these very abundant organic molecules occurs in a variety of habitats under either aerobic or anaerobic conditions and organisms capable of utilizing hydrocarbons as a sole energy source are present in almost every environment. The ability of the microbiota in a given environment to degrade the hydrocarbon pollutants present is chiefly dependent on the structure of the hydrocarbon compounds and on oxygen availability, as both of these variables determine which pathways are required for degradation of that specific structure. The degradation efficiency, however, is strongly dependent on various environmental and physical factors, for example temperature, pH, amount of nutrients, and bioavailability of the hydrocarbons. Many bacteria can use hydrocarbons as an energy source, but most of the time, that source of energy is not the preferred one, as compounds such as sugars or amino acids that are more readily convertible into substrates in the central energy metabolism pathways such as glycolysis or the tricarboxylic acid cycle are used in preference. However, hydrocarbonoclastic bacteria are highly specialized in utilizing hydrocarbons and play a key role in degrading hydrocarbons in polluted sites [14, 15].

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9.2.1 Aerobic degradation Oxygenase-mediated hydrocarbon degradation is quite commonly encountered in aerobic habitats such as surface soils, where it is carried out by a wide variety of bacteria, fungi, and algae. Aliphatic hydrocarbons (n-alkanes) are most commonly degraded through α- and ω-hydroxylation of the alkane to alcohol, catalyzed by alkane monooxygenase [16]. In pseudomonads, electrons for the reaction are relayed through an electron transport chain that involves rubredoxin and a FAD-dependent rubredoxin reductase [17]. Aerobic alkane metabolism is exemplified by the Pseudomonas oleovorans alkBFGHJKL operon, which encodes the enzymes required for conversion of alkanes to acetyl-coenzyme A [16]. The degradation pathways of alkenes and alkynes are highly similar to alkane-degrading pathways, but they can also undergo other additional reactions because of their unsaturated nature [18]. Degradation of aromatic hydrocarbons is similarly initiated by oxygenasemediated hydroxylation, resulting in the formation of catechols, followed by ring cleavage and aldehyde or carboxylic acid formation [16]. Typically, the initial reaction is mediated by a dioxygenase [19]. For polycyclic aromatics, naphthalene degradation by Pseudomonas putida serves as a well-studied model. It involves three plasmidborne operons, with one (nahAaAbAcAdBFCED) encoding the enzymes required for the conversion of naphthalene to salicylate, the second (nahGTHINLOMKJ) those for the conversion of salicylate to tricarboxylic acid cycle intermediates, and the third (nahR) encodes a regulator for the other two [16, 20].

9.2.2 Anaerobic degradation Due to the critical role of O2-dependent oxygenases in biodegradation of both aromatic and aliphatic hydrocarbons, it was generally assumed until about the 1990s that hydrocarbon biodegradation was an exclusively aerobic process. Multiple studies have since demonstrated the presence of hydrocarbon oxidation in anaerobic environments. Under anaerobic conditions, the O2-dependent oxygenase-catalyzed reactions are, for the most part, rendered irrelevant, although there are a few examples in the literature of intra-aerobic anaerobes capable of deriving oxygen from chlorate or nitrate and can thus employ monooxygenases for hydrocarbon degradation even in anaerobic environments [21, 22]. For the most part, however, anaerobic hydrocarbon oxidation requires alternative means of C-H bond activation through a variety of less well-known reactions using electron acceptors such as sulfate or nitrate [16, 23, 24]. A comparatively well-studied example is the oxidation of aromatics through the action of toluene-activating benzylsuccinate synthase and related glycyl radical-bearing alkyl or arylalkylsuccinate synthases. This enzyme adds toluene to a fumarate cosubstrate, forming a benzyl-substituted succinate [25, 26]. Among other examples

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are the O2-independent hydroxylation of ethylbenzene and ATP-dependent and ATPindependent dearomatization of benzoyl-CoA [26]. Anaerobic hydrocarbon oxidation by various members of Betaproteobacteria, Deltaproteobacteria, and Clostridia is now recognized to be a significant contributor to hydrocarbon turnover in nature, occurring wherever hydrocarbon loads exceed oxygen availability, such as in contaminated groundwater and other subsurface environments [27].

9.3 Effects of environmental conditions Since every environment is unique, the bioremediation process sometimes needs to be tailor-made for that environment. The remediation possibilities of polluted sites is determined by the composition of the hydrocarbons, the microbial community composition, and the environmental conditions [28]. For effective bioremediation, it is important to know the physical and chemical composition of the contaminated soil and the composition of the microbiota present because the soil composition, along with the microbiome composition, dictates the biochemistry predominant at the contaminated site. Thus, the choice and the design of bioremediation techniques have to be in accordance with the biochemical processes, the bioavailability, and bioactivity at the polluted site [29, 30]. However, the environmental factor is often confounded by various complications since it can be difficult to tamper with. In the context of the Arctic environment, the most obvious physicochemical variable to consider is temperature. Low temperature has a profound effect on biodegradation, not only in terms of biological activity, which is naturally hampered by lower reaction rates as dictated by basic thermodynamics, but it also affects various physicochemical properties of both the environment itself and the pollutant present [13]. Hydrocarbon degradation is therefore more problematic in cold environments because the viscosity of the pollutant is higher and the solubility is lower [31]. Because of this, the physicochemical features of the environment are of primary concern in bioremediation. So, the bottleneck of bioremediation is usually not the biological pathway potential of the bacteria, but rather limitation by the bioavailability and solubility of the targeted pollutant, which both can be adversely affected by lower temperature. Nevertheless, it has been demonstrated that microbial activity in Icelandic soil is governed by substrate availability rather that temperature [32, 33], supporting the viable idea of biostimulation in colder areas. Studies have shown that bioremediation on petroleum hydrocarbons with psychrotolerant bacteria can result in degradation of pollutants at 10°C, which is similar to the efficiency of bacteria at 30°C [34]. Freeze-thaw cycles in polar soil have even been reported to possibly stimulate the degradation of hydrocarbons in arctic soil [35, 36]. Therefore, the use of autochthonous microbial community is often the favored approach, since these microbes are already adapted to the cold environment and fluctuations in temperature. Putting the indigenous bacterial population to work through intrinsic bioremediation or biostimulation has been suggested to be the most feasible way of remediating hydrocarbons from cold environment [37].

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Intrinsic bioremediation is the process of letting the environment “heal itself,” but observing and managing the indigenous microbial community while biodegrading the pollutant without enhancing the process. Biostimulation refers to the closely related techniques of when the indigenous microbial community is stimulated to overcome the barriers in metabolic pathways and the limitations of degradation. That can be attained by providing nutrients or other compounds that enhance the biodegradation in any way, like oxygen or other electron receptors. Arctic soil is usually limited in nitrogen as well as phosphorous, and fertilizing arctic soil with nitrogen can enhance the biodegradation of hydrocarbons tremendously [38, 39]. A study on sub-Antarctic soils showed that 95% of the total hydrocarbon pollution was degraded by indigenous microorganisms within a year at 0–7°C [40]. In that study, the use of fertilizer was more effective in stimulating the assemblage of hydrocarbondegrading bacteria in a desert Antarctic soil than in a vegetated soil. Overfertilizing can, however, result in low water activity and, therefore, inhibition of microbial hydrocarbon degradation [41]. Sanscartier et al. [42] studied the potential for on-site bioremediation and different hydrocarbon removal processes in a polar desert in high arctic soils, where the annual mean in temperature is -15°C. They suggest that in dry, cold deserts, remediation of low-molecular hydrocarbons (nC16). In that study, biostimulation with the addition of surfactant was found to be the most effective treatment of the hydrocarbon contaminated soil [42]. The bioavailability of the targeted chemicals is another very important feature regarding bioremediation of pollutants. The fact that the most major hydrocarbon pollutants are insoluble or poorly soluble in water makes it difficult for organisms to access the target chemical in order to degrade it. In that case, measures to alter the bioavailability of the pollutants may be important [4, 30]. In addition to supporting the mixing of waterinsoluble hydrocarbons, biosurfactants have been shown to slow down the dispersion of hydrocarbon pollutants during freeze-thaw cycles and by that favoring further bioremediation [43]. In fact, biosurfactants are thought to be a promising solution to limited bioavailability in contaminated environments due to the chemical structure of the pollutant or other physicochemical barriers of the environment.

9.4 Microbial life in cold deserts The environmental conditions in Arctic desert soils, such as in the Icelandic highlands, the Canadian high Arctic, or in glacial moraines and forefields in Svalbard and Greenland, are highly restrictive, characterized by year-round low temperatures, freeze-thaw cycles, low water retention, regolith-like “soil” and largely abiotic sediments, low organic carbon content, and high salinity, resulting in a near total lack of visible vegetation [44]. There are pronounced phylogenetic dissimilarities between bacterial communities in unvegetated soil versus vegetated developed soil [45], possibly because of the different roles they play in harvesting energy and nutrients.

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The microbial communities found in these desert environments are thus, to a large extent, borne by the rock-weathering actions of chemolithotrophic bacteria such as iron and sulfur oxidizers and by carbon and nitrogen fixation by cyanobacteria [46]. Indeed, Actinobacteria and Cyanobacteria are far more abundant in unvegetated soils than in vegetated developed soil (Zumsteg et al., 2012). Nevertheless, complex microbial communities have been described in these environments in a number of studies. For example, glacial drift sheets in the Darwin-Hatherton glacier region of Antarctica were found to harbor a community comprising members of eight phyla: Actinobacteria, Bacteroidetes, Deinococcus-Thermus, Proteobacteria, Gemmatimonadetes, Firmicutes, Verrucomicrobia, and Planctomycetes [47]. In the Arctic region, microbial desert communities tend to be even more complex than those in comparable alpine or Antarctic environments [48]. Nevertheless, a characteristic community structure can be elucidated, strongly dominated by Proteobacteria, Actinobacteria, and Chloroflexi [49], whereas the Acidobacteria found to be characteristic of Arctic tundras [50] are generally not found in large numbers in Arctic deserts [49]. In recent years, bacterial communities in glacial forefields have been studied in terms of understanding the bacterial communities in poor developed soil and bacterial community succession in the Arctic. Even though low cell number and low activity of bacterial cells can be found in young forefields, the diversity of the bacterial community in poor developed and desert soil is surprisingly high [51]. Proteobacteria, Actinobacteria, Acidobacteria, Firmicutes, and Cyanobacteria showed high diversity in arctic forefields, and the abundance of Alphaproteobacteria increases while the abundance of Betaproteobacteria decreases in unvegetated soils [52]. Indeed, even the ice caps themselves are home to complex communities, as evidenced by a comparative study of the microbial communities present in the Icelandic glaciers Snæfellsjökull, Langjökull, Eyafjallajökull, Vatnajökull, Drangajökull, and Hofsjökull. In this study, members of the classes Betaproteobacteria, Alphaproteobacteria, Sphingobacteria, and Saprospirae were found to be the most abundant [53]. Furthermore, a recent metagenomic study of Greenland cryoconite communities found surprising abundance of genes involved in the degradation of polyaromatic hydrocarbons and polychlorinated biphenyls, indicating that the accumulation of these pollutants in Greenland glacier cryoconite has impacted its microbiota composition [54]. There is thus clearly a diverse autochthonous biota present in cold desert and ice environments that can be expected to be affected by polluting input, resulting in increased natural degradation. Nevertheless, several limiting factors can also be expected to limit natural biodegradation of pollutants in these environments.

9.4.1 Bioprospecting cold desert soils for hydrocarbon-degrading microbes The Arctic environment has attracted considerable attention from bioprospectors in recent years [55–58]. Arctic soil microbiotas are recognized as valuable sources

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of cold-active enzymes, biosurfactants, and various metabolites, as well as for coldadapted bioremediation [59–63]. While the numbers of hydrocarbon degraders are usually low or undetectable in pristine Arctic soils, contaminated soils often display high numbers of hydrocarbon degraders that may persist for decades after the initial spillage [64]. Although only a small part of the resident microbiota can be expected to be easily culturable, the aerobic bacteria most readily utilizable as bioremediators can be expected to grow with relative ease in many commercially available growth media. It therefore seems feasible to prospect contaminated cold soils for biodegraders of alkanes and aromatics, and, indeed, a large number of such efforts can be found in the literature. Hydrocarbon-degrading members of the genera Rhodococcus, Pseudomonas, and Sphingomonas appear most often in these studies [64], although several other taxa have been reported, such as Pedobacter [65], Arthrobacter [66], Acidovorax, and Variovorax [67], among others. An obvious requirement for microorganisms to be used for in situ bioremediation in Arctic environments is that it be cold-adapted. While it seems reasonable to expect isolates from cold deserts to be adapted to the cold, it is important to remember that summertime temperatures in surface soils may periodically approach 20°C, and thus, some of the bacteria thriving in that environment will have optimal growth rates above the psychrophilic range (≤15°C). Nevertheless, many of the hydrocarbondegrading bacteria isolated from cold desert environments have indeed been shown be psychrotrophic and oxidize hydrocarbons at low temperatures. For example, Sphingomonas Ant 17 degraded phenanthrene at 4°C, although higher degradation rates were observed at 28°C [68], and Rhodococcus Q15 mineralized alkanes at 0°C [69], which is traditionally considered to be at the lower temperature threshold for significant hydrocarbon biodegradation, meaning that bioremediation in terrestrial polar environments is likely limited to the warmer summer season [13]. However, more recent work has shown this traditional view to be too simplistic as wintertime bacterial activity under snow cover, often at subzero temperatures, has been found to be an important factor in Arctic ecology [70, 71]. It should be borne in mind, however, that there are more limiting factors to hydrocarbon degradation rates in cold climates than simply the microorganism’s psychrotolerance. As discussed above, temperature will also affect the physical properties of petrochemicals. At lower temperature, the viscosity of oil is increased, the volatilization of short-chain alkanes is reduced, and water solubility decreased. The bioavailability of hydrocarbons is thus generally decreased at lower temperatures [72]. Nevertheless, hydrocarbon degradation has been demonstrated in situ to occur in soils at subzero temperatures in Svalbard [73]. Tolerance of wide temperature fluctuations and freeze-thaw cycles are also among requirements for potential bioremediators in Arctic soils. Indeed, some of the hydrocarbon-degrading isolates from cold soils have been found to be well adapted to freeze-thaw cycles. For example, Sphingomonas Ant 17 was found to be more tolerant to freeze-thaw cycles than was the mesophilic Sphingomonas WPO-1 [68] and freeze-thaw microcosm experiments in the presence of diesel fuel indicated that the

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freeze-thaw regime led to the predominance of hydrocarbon-degrading rhodococci in the microcosms [35]. Similarly, a study of hydrocarbon-contaminated Resolution Island-soil communities and their responses to freezing-thawing cycles showed an increase in 14C-hexadecane mineralization and the emergence of Corynebacterineae (of which Rhodococcus is a member) during the freezing phase [36].

9.5 Concluding remarks Although the Arctic deserts and glacial moraines appear barren to the casual observer, they are in fact home to a diverse, abundant, and active microbiota, within which easily culturable hydrocarbon-degrading bacteria can be found. These organisms can be expected to be adapted to harsh Arctic conditions, including low temperature, freezethaw cycles, and low organic carbon content and may thus be utilizable for partial bioremediation of contaminated sites in the delicate Arctic environment, although large-scale testing of their applicability to real-world situations is mostly lacking. The autochthonous microbiota of Arctic desert and ice environments has also been shown to respond to pollutant input, suggesting that in situ biostimulation techniques may be considered when contemplating bioremediation in these environments.

References [1] Atlas RM. Microbial bioremediation in polar environments: current status and future directions. In: Bej AK, Aislabie J, Atlas RM, editors. Polar microbiology: the ecology, biodiversity and bioremediation potential of microorganisms in extremely cold environments. Boca Raton, FL: Taylor & Francis; 2009. p. 373–91. [2] Gunnarsdóttir R, Jenssen PD, Erland Jensen P, Villumsen A, Kallenborn R. A review of wastewater handling in the Arctic with special reference to pharmaceuticals and personal care products (PPCPs) and microbial pollution. Ecol Eng [Internet]. 2013 Jan;50:76–85. Available from: http:// linkinghub.elsevier.com/retrieve/pii/S0925857412001528. Cited July 18, 2017. [3] Tyagi M, da Fonseca MMR, de Carvalho CCCR. Bioaugmentation and biostimulation strategies to improve the effectiveness of bioremediation processes. Biodegradation. 2011;22(2):231–41. [4] Speight JG, Arjoon KK. Bioremediation of petroleum and petroleum products. 2012. p. 1–567. [5] Singh P, Jain R, Srivastava N, et al. Current and emerging trends in bioremediation of petrochemical waste: a review. Crit Rev Environ Sci Technol [Internet]. 2017;47:155–201. Available from: http:// dx.doi.org/10.1080/10643389.2017.1318616. [6] Herrero M, Stuckey DC. Bioaugmentation and its application in wastewater treatment: a review. Chemosphere [Internet]. 2015 Dec;140:119–28. Available from: http://linkinghub.elsevier.com/ retrieve/pii/S0045653514012181. Cited June 30, 2017. [7] Trujillo ME, Kroppenstedt RM, Fernandez-Molinero C, Schumann P, Martinez-Molina E. Micromonospora lupini sp nov and Micromonospora saelicesensis sp nov., isolated from root nodules of Lupinus angustifolius. Int J Syst Evol Microbiol. 2007;57:2799–804. [8] Fernández-Luqueño F, López-Valdez F, Sarabia-Castillo CR, García-Mayagoitia S, Pérez-Ríos SR. Bioremediation of polycyclic aromatic hydrocarbons-polluted soils at laboratory and field

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Helga Stan-Lotter

10 Subsurface extremophiles and nuclear waste storage Abstract: Deep geological disposal is considered by many countries the best way of managing nuclear waste. Rock salt, granite, claystone, and other geological formations are already in use or being prepared for storage of radionuclides. Underground rock laboratories have been established for research purposes. Geologically old rock salt is a particularly suitable medium for permanent waste isolation and has a good record of performance in the USA. Issues arising from the presence of subsurface microbial life and storage of radioactive waste are reviewed here. Liquid forms of radioactive waste are the greatest portion of materials for disposal. Efforts for concentration of liquids include methods for adsorption of uranium and other radionuclides to microbial cells or cellular components. Another promising method is dissimilatory reduction of radionuclides by microorganisms, which increases the insolubility of nuclear waste and thus renders it more suitable for long-term storage. The capacities for remediation purposes of extremely halophilic bacteria and archaea, which are indigenous to rock salt, are just beginning to be explored. Nuclear transmutation – the fission of radionuclides by irradiation with fast neutrons into less harmful elements – is increasingly being developed by several institutions; however, the need for geological waste disposal will most certainly remain for many years.

10.1 Introduction Salt mines have gained some notoriety as storage rooms for precious works of art, gold reserves, and foreign currency, notably during World War II, in Germany and Austria (Fig. 10.1). The mines provided shelter from bombing and the vicissitudes of weather above ground, and their interior provides stable environmental conditions. Salt mines are relatively dry, with a low average relative humidity of 40%. There is very limited exposure to ultraviolet light. The temperature in depths between 600 and 800 meters ranges between 7°C and 10°C. Fluctuations of temperature and humidity are nearly absent. This stable environment matches in some ways the preferred climate-controlled conditions for document storage today (from [1]). The time frame for the purpose of saving precious materials was rather short, in the range of weeks or perhaps months, certainly not beyond a human lifetime.

https://doi.org/10.1515/9783110424331-010

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Fig. 10.1: The US generals Dwight D. Eisenhower, George S. Patton Jr., and Omar N. Bradley (from right to left) on April 12, 1945, inspect paintings from museums in Berlin, which were stored by Nazis in the salt mine of Merkers, Thuringia, Germany (File: Eisenhower, Bradley and Patton inspect looted art HD-SN-99-02758.JPEG – Wikipedia; public domain).

Radionuclides and other nuclear waste have to be stored safely on quite a different time scale. Radioactive waste is a by-product of nuclear power generation and other applications of nuclear fission or nuclear technology, such as research and medicine. The radioactivity decays with time, and thus, radioactive waste has to be confined in appropriate disposal facilities for a sufficient period until it no longer poses a threat. The storage time of radioactive waste depends on the types of waste and radioactive isotopes. It can range from a few days for shortlived isotopes to millions of years. For example, the spent fuel rods from nuclear reactors contain actinides that emit alpha particles, such as 234Uranium (half-life 2.45 × 105 years), 237Neptunium (half-life 2.14 × 106 years), 238Plutonium (half-life 87.7 years), and 241Americium (half-life 433 years) [2]. Current major approaches to managing radioactive waste have been segregation and storage for short-lived waste, near-surface disposal for low- and some intermediate-level waste, and deep burial for the high-level waste. A brief note on the “classification” of radioactive wastes is appropriate here (see [3]). The distinctions between low-level, intermediate-level, and high-level waste categories (LLW, ILW, HLW, respectively) are not consistent internationally. Classification schemes may be based on half-life, activity, origin or source, degree of isolation required, etc. In general, low-level waste contains radionuclides with low activities and short half-lives and generates no heat.

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Intermediate waste may contain radionuclides with low to intermediate activities and short to long half-lives, generating no to negligible heat. High-level waste contains radionuclides with high activities and long or short half-lives or both and generates heat. Besides deep salt deposits, which are thought by some agencies, scientists, and administrations to be particularly suitable, as is discussed in this article, disposal concepts have been developed for shale, volcanic rock, granite, and old mines (see Tab. 10.1). Microbial activity can play a major role in the performance of geological radioactive waste repositories. The notion of a “deep hot biosphere” was suggested in 1992 by Thomas Gold [4], whose paper became a stimulus for much of today’s deep biology search. Gold [4] had estimated that if all life inside the Earth were brought to the surface from as much as 5 km down, it might form a layer 1.5 m thick, covering all land surfaces and weighing more than all flora and fauna together. Some scientists believe that, indeed, subsurface life may be the main stage for the planet’s biodiversity [5], even if newer assessments have challenged the magnitude of the proposed biomass [6, 7]. In any case, numerous studies showed that microbial effects on the migration or retardation of heavy metals, including radionuclides, as a result of metal-binding and redox mechanisms can occur, as well as the production of gas and various organic molecules. These are significant events, both for sealed geologic radioactive waste repositories and for prokaryotes in contact with radionuclide containing materials in the environment. Numerous species of bacteria and archaea can tolerate the extreme environments of radiation, desiccation, alkalinity, acidity, heat, high ionic strength, and pressure, which are to be found in geological repositories. In addition, microbial populations are diverse and can be unique to their individual environments, and therefore, microbial effects should be considered on a per-site or case-by-case basis. Another aspect of microorganisms and radionuclides is concentration and remediation efforts. For example, liquid radioactive wastes were generated at the Savannah River Site, USA, as by-products from the processing of nuclear materials for national defense, research, and medical programs. The waste, totaling about 36 million gallons (about 140 million liters), is currently stored in 44 underground carbon-steel waste tanks [8]. This radionuclide-contaminated water can migrate into surface waters, where it becomes a threat to organisms and water supplies. Other liquid wastes are produced in nuclear power plants, medicinal procedures, and in research, which lead to additional disposal issues. It is desirable to concentrate or filter these types of waste, to arrive at smaller volumes of radionuclides for easier storage, and methods for biosorption of radionuclides in liquids with the aid of microorganisms have been investigated [9, 10]. This chapter describes some aspects of major underground storage sites for radioactive waste and underground rock laboratories (URLs), lists recognized problems originating from subsurface microorganisms, and presents biotechnological developments for bioremediation of radionuclides.

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10.2 Subsurface storage and research sites 10.2.1 Deep geological facilities In the scientific community worldwide, it is accepted that deep geological disposal is the appropriate way to dispose of spent or used fuel elements, high-level radioactive waste, and other, especially long-lived radioactive waste safely and securely for long times. Several types of geological formations were considered to host deep geological repositories and include argillaceous rocks, crystalline rocks, and rock salt [11]. Underground spaces, whether large, individually engineered cavities or the small but numerous natural voids in reservoir rocks, have temporarily or permanently accommodated water, brine, liquid and gaseous hydrocarbons, compressed air, and dangerous materials [3]. Excavations in low-permeability crystalline basement rocks, e.g. gneiss and granite, are currently being used to dispose of some categories of radioactive waste. Former limestone and uranium mines are serving the same purpose. Thick, geologically old rock salt is another confinement medium suitable for permanent waste isolation [3]. Tab. 10.1 shows an overview of deep geological storage sites that have been or still are in use, as well as some planned facilities. Until permanent storage sites are constructed, used nuclear fuel and other wastes have to be stored in interim storage facilities. It can be mentioned here that public concerns about the safety and environmental issues of subsurface radioactive waste storage have often led to postponements or possibly cancellations of the development of new sites (e.g. in Germany, Gorleben salt deposit). Tab. 10.1: Deep geologic repositories for radioactive waste in use or under construction. Country

Name

Geology (host rock)

Depth

Status/Operation

Reference

Canada

OPG DGR (LLW, ILW)

Limestone

680 m

License application in progress 2017

Czech Republic

Hostim

Limestone

30 m

1959–1965

[3]

Czech Republic

Bratstvi

Uranium mine

n/i

Since 1974

[3]

Finland

Olkiluoto

Tonalite

60–100 m

Since 1992

[16]

Finland

Loviisa

Granite

120 m

Since 1998

[16]

Germany

Gorleben

Salt dome

850 m

Proposed, on hold

[3]

Germany

Asse

Salt/potash mine

750 m

1978–1998

[3]

[15]

10.2 Subsurface storage and research sites 

 247

Tab. 10.1 (continued) Country

Name

Geology (host rock)

Depth

Status/Operation

Germany

Schacht Konrad (shaft repository)

Iron ore mine 800–1300 m Under construction

Russia

Three locations: LLW ILW HLW

Clastic sediment

Sweden

Forsmark (LLW and ILW)

Granite

50 m

Sweden

Forsmark (spent fuel)

Granite

450 m

Reference [12]

Since 1963

[3]

Since 1988

[13]

License application 2011

[17]

180 m 355 m 500 m

LLW, low-level waste; ILW, intermediate-level waste; HLW, high-level waste; n/i, no information

The Swedish Final Repository (SFR) for short-lived radioactive waste in Forsmark started operating in 1988 and was then the first of its kind in the world. The radioactive waste deposited there is low- and medium-level waste. This means that unlike spent nuclear fuel, it does not have to be cooled and is relatively short-lived. The repository is situated 50 m below the bottom of the Baltic Sea [13]. Most of the waste deposited in the SFR comes from the operations of Swedish nuclear power plants. It can include filters that have collected radioactive substances from reactor water, tools, and protective clothing. But radioactive waste from hospitals, veterinary medicine, research, and industry is also deposited [13]. A repository for high-level waste in 500 m depth is planned. Pedersen [14] describes the deposition tunnels and the foreseen encapsulation of spent fuel cells in copper canisters, as well as their shielding from groundwater by bentonite clay.

10.2.2 Underground rock laboratories For research and testing purposes, URLs in granitic rock have been built in Canada (Whiteshell URL, closed in 2010), Finland (ONKALO URL), Sweden (Äspö URL in 500 m depth), and Switzerland (the Grimsel test site), while another one is under construction in Japan (Mizunami URL) [18]. Of interest is also the Boulby International Subsurface Astrobiology Laboratory, UK, in a depth of 1100 m [19], which is located in a salt mine. Because these URLs are research facilities, hydrological and geochemical sampling programs are extensive and they generate a wealth of background information about the sites. Since the quest to build HLW repositories that are safe

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for several hundreds of future generations takes a long time, many of the research programs will last for decades. This is the case for microbiological investigations in the URLs of Finland and Sweden, where microbiology has been investigated since over two decades [18]. Since microorganisms are regarded to be a concern in most or perhaps all types of geological storage sites, potential effects of the presence of native and/or introduced microorganisms are an important part of research in all URLs.

10.2.3 Rock salt deposits Since many years, countries like the United States of America, the Federal Republic of Germany, The Netherlands, Poland, and recently, the Czech Republic, consider rock salt as one of the favorable rock types to host a deep geological repository for different kinds of radioactive waste, including high-level waste and spent nuclear fuel. An advanced scientific and geotechnical understanding of rock salt as an appropriate geological material to host repositories has been accumulated by the dedicated research carried out by a number of countries favoring this material. The most important positive attributes of rock salt for use as a repository host rock are considered to be [11]: –– extremely low permeability that isolates the waste from any near surface groundwaters; –– dry environment; –– high thermal conductivity; –– viscoplastic behavior that increases with temperature and pressure and closes all void spaces; –– predictable geology; –– a vast store of knowledge and experience on excavation techniques, mine development, and safe operation is available, stemming from salt mining at great depth; –– rock salt is easy and inexpensive to excavate; –– transparent conceptual and numerical models (enhances scientific and public acceptance) are at hand, and –– theoretical and operational waste experience bases are readily available (reduces the learning curve and cost for new projects). Judging mainly from German and US data, rock salt is a very promising geologic medium for the development and safe as well as cost-effective operation of underground research laboratories and repositories for long-lived and heat generating radioactive waste [11].

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10.2.4 Advantages of the Waste Isolation Pilot Plant The Waste Isolation Pilot Plant (WIPP) near Carlsbad in southeastern New Mexico started in 1999 to dispose of intermediate-level (transuranic) radioactive waste contaminated with radionuclides that have long half-lives [3]. The repository horizon is located at a depth of 655 m in a Permian salt deposit called the Salado formation [20]. As far back as 1957, the National Academy of Sciences, USA, has recommended salt for radioactive waste disposal because at depth, it would plastically deform, a motion called “salt creep” in the salt-mining industry, to close and seal any openings created by mining, and also in and around the waste [20]. Indeed, to test the isolation capability of underground storage from the accessible environment, dramatic detonation experiments were carried out by the USA [2]. In salt formations, three nuclear devices were detonated, one in the Salado formation near the present WIPP facility in New Mexico and two others at the Salmon Site in Mississippi. The shot near the WIPP site had a magnitude of 3.1 kilotons, which is in the range of underground nuclear explosions detectable by seismic signals. By virtue of monitoring results, the radioisotopes were determined to be confined to the test cavity; i.e. the test cavity was determined not to be leaking any radioactivity [21]. No migration has been since detected outside the experiment’s boundary for half a century. It was concluded that salt formations have been shown to seal and confine nuclear detonations [2, 3]. A major intrinsic advantage of a salt repository is the lack of groundwater to seal against. Even though regional aquifers may be close to the host unit, the sealing systems can be designed to perform in contact with groundwater [2]. If water flow occurs within the repository openings, the chemistry of water or brine could impact engineered materials. However, the materials used – e.g. concrete, asphalt, clay – are durable, with minimal potential for degradation or alteration [2]. Microbial degradation, material interactions, and mineral transformations are often incompletely understood and therefore continue to be the focus of ongoing research (see below). Degradation of concrete is possible, but unlikely as only small volumes of groundwater will ever reach the concrete [2]. No comments were made on potential microbial degradation of bentonite by [2], but microbes were shown to survive in compacted bentonite clay [14, 22], provided enough moisture was available. Fig. 10.2 shows containers with radioactive waste in an underground storage room at WIPP. Operations utilized at WIPP include stacking of contact-handled waste on the floor and horizontal disposal of remotely handled waste in pillars as shown in the photograph.

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Fig. 10.2: Disposal operations for transuranic (TRU) waste at the Waste Isolation Pilot Plant (WIPP). Waste containers are either contact-handled, as those on the floor, or remotely handled as that on the left side (from [2]; unlimited release).

Sealing a nuclear waste repository and waste corrosion are considered two primary issues with salt disposal. The Sandia report from 2011 [2] points out historic analogues from mining as follows: Anthropogenic evidence associated with vast and pertinent mining experience provides important qualitative assessments of preserved artifacts. The Hallstatt area in Austria supported prehistoric salt mining. Archeological reexcavation has recovered organic material such as leather, wood, clothing, and even an unfortunate Celtic miner preserved in the salt for 3000 years. More recent analogues spanning 50 or 100 years of mining experience offer convincing evidence of structural, mechanical, and hydrological behavior of the underground salt environment. The salt and potash mining industry has placed functional seals in underground workings, backfilled, and reconsolidated crushed salt in active mining operations. Within the potash basin in southeastern New Mexico mining operations, machinery that is not brought back to surface suffered almost no corrosion after more than 50 years.

10.3 Microorganisms and radionuclide storage As pointed out in the Introduction, subsurface environments are populated by various microorganisms, and the effects of microbial activity in geological repositories for nuclear waste are an intensive area of research. The Atomic Energy of Canada Limited (AECL) initiated studies to address the potential of microbial activities on the integrity of the multiple barrier system for nuclear fuel waste disposal as early as 1982, and the results of research up to 1997 are reviewed in [23]. The main issues of that microbial

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program are still of great interest and importance and are stated here briefly (modified from [23]): –– Survival of microorganisms in buffer materials under relevant radiation, temperature, and moisture conditions; –– Potential for microbially influenced corrosion (MIC) of container materials; –– Transport of microorganisms through highly compacted buffer material in relation to MIC and radionuclide migration; –– Introduction of microbial nutrients through human presence and activity during vault construction and operation; –– Potential for microbial gas production in backfill materials; –– Presence and activity of microorganisms in deep underground waters; –– Effects of biofilms on radionuclide migration in the geosphere. The Sandia National Laboratory in its 2011 report [2] has accumulated a list called Features, Events, and Processes (FEPs), which are to be used in the certification and recertification of the Waste Isolation Pilot Plant or other sites to be considered as repositories. In the FEPs numbering scheme for WIPP, FEPs are categorized as “Natural FEPs,” “Wasteand Repository-Induced FEPs,” and “Human-Induced Events and Processes.” This list provides a valuable resource as a starting point if the USA chooses a salt option for disposal of high-level nuclear waste. Some of the waste and repository-induced FEPs in the list (designated with W) are concerned with microbial activities and are shown here: W44 – Degradation of Organic Material – Microbial breakdown of cellulosic material in the waste will generate gas, W45 – Effects of Temperature on Microbial Gas Generation – Temperature rises could affect the rate of microbial gas generation. W46 – Effects of Pressure on Microbial Gas Generation – Increases in gas pressure could affect microbial populations and gas generation rates. W47 – Effects of Radiation on Microbial Gas Generation – Radiation could affect microbial populations and, therefore, gas generation rates. W48 – Effects of Biofilms on Microbial Gas Generation – Biofilms serve to maintain optimum conditions for microbial populations and affect gas generation rates. W76 – Microbial Growth on Concrete – Acids produced by microbes could accelerate concrete seal degradation. W87 – Microbial Transport – Radionuclides may be bound to or contained in microbes transported in groundwaters. W88 – Biofilms – Biofilms may retard microbes and affect transport of radionuclides.

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10.4 Removal of uranium and other radionuclides Liquid radioactive wastes constitute by far the greatest bulk of radioactive wastes produced by various industries. Liquid radioactive waste was initially dumped in freshwater or in marine reservoirs and in underground storage [24]. Major disposal of liquid wastes underground occurred in the former Soviet Union and the USA [8, 9, 24]. In several cases, storage has been compromised, leading to contamination of trillions of hectoliters of groundwater and millions of cubic meters of contaminated soil and debris [9], with a probably astronomical budget needed for the costs of cleaning up these sites.

10.4.1 Adsorption to free or immobilized cells Several working groups investigated microorganisms for the removal of uranium from liquid environments. Biosorption – the metabolism-independent sorption of heavy metals and radionuclides [8] – was taking place, with uranyl ions attaching to cellular components, most often, but not exclusively, to the cell walls. The carboxyl groups of the bacterial peptidoglycan are the main binding site for cations in Gram-positive cell walls, while phosphate groups are binding sites in Gram-negative species [25]. Chitin, a polymer of N-acetylglucosamine, is a component of fungal cell walls and acts as effective biosorbent for radionuclides, as do chitosan and other chitin derivatives [25]. Inactivated or killed cells were equally or even more efficient in their capacity to adsorb to uranium ions in liquid environments. In other procedures, cellular metabolism was essential for the accumulation of uranium, since reduction of usually U (VI) to U (IV) was occurring, which rendered the uranium less soluble or, depending on the ionic environment, even insoluble (see below). Following are descriptions of several variants of bioremediation by biosorption using various microorganisms, except extreme halophiles, which were isolated from rock salt. These are treated in Section 4.3. Nakajima et al. [26] recovered uranium from sea and fresh water using immobilized Streptomyces viridochromogenes and Chlorella regularis cells. The cells that had been immobilized in polyacrylamide gel were much more stable than free cells; they were not susceptible to microbial degradation and are therefore of potential interest for remediation processes on an industrial scale. Adsorption of uranium was reverted easily by treatment with a solution of Na2CO3, thus allowing repeated use in the adsorption-desorption process. DiSpirito et al. [27] reported on the uptake and cellular distribution of UO22+ by washed cell suspensions of Thiobacillus ferrooxidans. Accumulation of cellassociated uranium was determined by collecting cells on membrane filters. Cells that had been inactivated by either ultraviolet radiation or potassium cyanide accumulated about 40% more uranium than did viable cells. Most of the uranium was associated with the cell wall and membrane fractions, and relatively little uranium

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was detected in the cytoplasmic, lipopolysaccharide, and periplasmic space material fractions. Galun et al. [28] were the first workers to show biosorption of uranium to fungal cells (Penicillium digitatum). Here, killing of the fungal biomass increased the uptake of uranium, depending on the solvent or denaturant used, by up to 2.5-fold compared to the living biomass to about 31 ug uranium per gram of dry mycelium. These agents were thought to expose uranium binding sites either by configurational change or by removal of masking groups. Tsuruta [29] tested the removal and recovery of uranium using microorganisms, which had been isolated from Japanese uranium deposits. Uranium was supplied as UO2(NO3)2, or in waste water from uranium refining processes, or from mine tailings. The capacity of different strains adsorbing uranium at the optimum pH of 6.0 varied between 10% and 98%. The highest capacity was found in a Lactobacillus strain, which could remove 2.3 micromol of uranium (547 ug) per gram dry weight of microbial cells. Desorption with EDTA showed that 93% of the uranium was in the cell wall and 7% was in the intracellular fraction. Sasaki et al. [30] described a novel combination of several previously applied laboratory protocols for the adsorption of mainly cesium radionuclides from polluted soil around the Fukushima Electric Power Plant. A photosynthetic bacterium, Rhodobacter sphaeroides SSI, was used, which was immobilized in alginate beads. The beads were put in a mesh bag and soaked in a suspension of 5 kg of soil in water, which resulted in a decrease in radioactivity by 31% after 15 days of aerobic treatment. With an additional pretreatment by anaerobic digestion and lactic acid fermentation with a culture of Lactobacillus casei, the radioactivity of the original soil had decreased by 67% within a total of 19 days. After treatment, the immobilized beads containing the radioactivity were incinerated, achieving 97–99% reduction in biomass volume and weight, without release of radioactive Cs into the air.

10.4.2 Dissimilatory reduction which decreases solubility Whereas certain of the just described procedures could in principle be performed like a chemical reaction, using dead microorganism or their extracted cellular polymers, for the following technologies, metabolically active microorganisms are required. The reduction of soluble U(VI) to less soluble U(IV) species was initially believed to be entirely abiotic in nature, with organic and inorganic compounds acting as reducing agents. This notion changed in the early 1990s when biotic uranium reduction was described for a dissimilatory Fe(III)-reducing bacterium [31, 32]. Since then, numerous bacteria have been found that are capable of reducing uranium. Of these bacteria, only Anaeromyxobacter dehalogenans, Carboxydothermus ferrireducens, Desulfotomaculum reducens, Geobacter metallireducens, and Shewanella putrefaciens have been reported to grow using U(VI) as a terminal electron acceptor [32].

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Gadd [25] reviewed dissimilatory biological metal reduction by microorganisms and pointed out the potential applications for uranium precipitation and its removal from solution, as well as the concentration of uranium from low-grade sources. As mentioned above, the basis of the process is the solubility of U(VI) compounds, in contrast to U(IV) compounds, such as the hydroxide or carbonate, which have low solubility and readily form precipitates at neutral pH. These precipitates may accumulate as a dark matter in association with the bacterial cells [25, 31]. Sulfate-reducing bacteria can also reduce U(VI) to U(IV) and it has been suggested that this reaction is responsible for observed high local concentrations of precipitated uranium in sediments under sulfate-reducing conditions [31]. Since bacterial U(VI) reduction can result in a very pure precipitate of U(IV) carbonate, it has been regarded as a viable alternative to uranium removal from aqueous solution by more conventional chemical treatments [25, 33]. Prakash et al. [34] reviewed the currently known microbially mediated mechanisms for biotransformation of radionuclides under various environmental conditions. Especially the biochemical basis such as enzymatic mechanisms and also recent developments in “-omics”-based technologies, i.e. genomics, transcriptomics, and proteomics, were treated. The first demonstration of dissimilatory U(VI) reduction by Lovley and coworkers [31] showed that the Fe(III)-reducing bacteria Geobacter metallireducens and Shewanella oneidensis (formerly Alteromonas putrefaciens and then Shewanella putrefaciens) can conserve energy for anaerobic growth via the reduction of U(VI). Other organisms including the sulfate-reducing bacteria Desulfovibrio desulfuricans [34] and Desulfovibrio vulgaris [35] also reduce uranium but are unable to conserve energy for growth via this transformation. 99 Technetium is another long-lived radionuclide (half-life 2.13 × 105 years) that mostly occurs in nuclear wastes. Tc(VII) has weak ligand-complexing capabilities and is difficult to remove from solution using conventional chemical approaches. Several reduced forms of the radionuclide are insoluble, however, and metal-reducing microorganisms can reduce Tc(VII) and precipitate the radionuclide [36]. Lloyd and Macaskie [37] were the first to demonstrate direct enzymatic reduction of Tc(VII) by microorganisms. X-ray absorption spectroscopy studies have recently identified insoluble Tc(IV) as the final oxidation state produced when Tc(VII) is reduced enzymatically by Geobacter sulfurreducens [38], Escherichia coli [36], and Shewanella putrefaciens [39]. Besides 238U and 99Tc, which are the highest-priority radionuclide pollutants in most medium- and low-level radioactive wastes, other actinides, including 230Th, 237 Np, 241Pu, and 241Am, occur at contaminated sites [40, 41]. Iron-reducing bacteria such as Geobacter sp, and Rhodoferax ferrireducens have the metabolic potential to reduce these radionuclides enzymatically [42]. Prakash et al. [34] pointed out that although it is possible for Fe(III)-reducing bacteria to reduce and precipitate actinides in one step [e.g. soluble U(VI) to insoluble U(IV)], few studies support the direct formation of an insoluble mineral stage; rather they indicate the formation of a cation prone to bioprecipitation [38]. This phenomenon

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is illustrated when considering highly soluble Np(V), which was reduced to insoluble Np(IV) by Shewanella putrefaciens, then removed as an insoluble phosphate biomineral by a phosphate-liberating Citrobacter sp. The studies by Lloyd and colleagues [38] have also suggested that the reduction of Pu(IV) to Pu(III) can be achieved by Fe(III)reducing bacteria, and Pu(III) was reported to reoxidize spontaneously [43].

10.4.3 Halophilic microorganisms and radionuclides Ancient salt deposits such as the Permian Salado formation are subsurface environments from which extremely halophilic microorganisms have been isolated, which belong predominantly to the archaea or haloarchaea [44–46]. The interactions of extremely halophilic microorganisms in deep salt repositories like the WIPP site with radionuclides are of interest, but few studies have been performed so far. Ams et al. [47] evaluated the adsorption of neptunium (V) at high ionic strength on Chromohalobacter sp., a halophilic bacterium, which was isolated from a briny groundwater near the WIPP site. In contrast to what is generally observed under relatively low ionic strength conditions, adsorption was shown to increase by as much as 25%, with increasing ionic strength from 2 to 4 M [47]. In the Helmholtz Research Center Dresden, Germany, uranium adsorption to halophiles from rock salt is currently studied [48]. Two strains of the extremely halophilic Halobacterium noricense are apparently indigenous to geographically very distant Permian salt deposits. The type strain Hbt. noricense DSM 15987 was obtained from a Permian salt deposit in the Austrian alps [49]. A very similar strain was isolated from a mixed culture of archaea enriched aerobically from WIPP halite [48]. Uranium (VI) was strongly bound to these haloarchaea, probably causing accumulations of cells and the formation of biofilm-like structures. This adsorption would decrease the mobility of radionuclides and could be beneficial for the safe performance of a repository. However, in the presence of chelating agents like citrate or EDTA, uranium is released and the haloarchaea do not provide an effective retention of the radionuclide [48]. A more detailed study of the molecular interactions between Hbt. noricense and uranium was presented in [50, 51]. Following incubation of cells of Hbt. noricense with uranium salts, uranium (VI) phosphates and carboxylates were formed, probably by reactions with DNA and other cell components, and adsorption of uranium compounds on the cell surfaces was observed. High- resolution microscopy images showed that cell morphology was preserved during the experiments. A partial reduction of uranium (VI) to uranium (IV) mediated by the cells was also noticed. Such pathways would lead from soluble uranium to insoluble phases, as described above (4.2), and could benefit the immobilization and/or retardation of uranium [50]. A comparison of Hbt. noricense with Brachybacterium sp. G1, a moderately halophilic isolate from the Gorleben salt mine in Germany, from a depth of 840 m, with

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respect to uranium bio-association was performed [51]. The adsorption of uranium to Brachybacterium sp. G1 was faster and generally higher, related to biomass, than that of Hbt. noricense. The results were explained with a higher number of carboxyl groups in Brachybacterium sp. G1, due to its peptidoglycan cell wall [51]. The migration behavior of uranium in rock salt and brines is thus definitely influenced by the presence and types of microorganisms. Swanson et al. [52], who investigated the biodegradation of actinide-complexing chemicals by indigenous halophilic microorganisms in the WIPP site, pointed out the similarities in organisms and DNA sequences from locations both geographically distant and temporally disparate, such as Permian salt deposits. A comparison was made with previous findings of similar strains of Halococcus salifodinae in different European salt deposits of Permo-Triassic age [53] and for strains of Halobacterium spp. [54]. Therefore, there is reason to believe that the organisms enriched from WIPP halite may also be present in the halites of other salt-based repositories and may play a role in the degradation of complexing agents for actinides in those as well [52].

10.5 Nuclear transmutation Finally, a different approach to radionuclide waste should be mentioned here, although no microorganisms are involved. Nuclear transmutation is the conversion of one chemical element or isotope into another. When irradiated with fast neutrons in a nuclear reactor, these isotopes can be made to undergo nuclear fission, destroying the original isotope and producing a spectrum of radioactive and nonradioactive fission products. The long-lived radioactive elements in waste are in this way converted to shorter-lived particles that produce radiation for a much shorter period and are less radiotoxic.  There have been proposals for reactors that consume nuclear waste and transmute it to other, less-harmful nuclear waste. Transuranium elements such as the isotopes of plutonium, neptunium, americium, and curium, which are radioactive for tens of thousands of years, can be transmuted to others that are radioactive for only hundreds or even tens of years. In some cases, the end products are stable, that is nonradioactive, isotopes. However, the OECD/Nuclear Energy Agency warns that these treatments should not be considered as an alternative to deep geological disposal and should not be presented as such [55]. In any case, procedures are under development, and even the most optimistic scenarios are forecasting several decades until the methods develop from laboratory scale to industrial size reactors [55]. Two examples of ongoing research are presented here (from [56]): 1. Transmutation experiments were carried out on 241Americium in the High Flux Reactor at Petten (Netherlands) in a project designed to study the “Experimental Feasibility of Targets for Transmutation” (EFTTRA). It was found that after

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irradiation for 350 days, 96% of the 241Americium was transmuted. However, the target pellets were swollen (by about 18%) due to helium release. This showed that modifications in the design of pellet fabrication were required. 2. An experiment carried out at the European Nuclear Research Centre (CERN) directly verified a concept of so-called resonance crossing for significantly enhancing the neutron-capture efficiency for long-lived fission products. In lead, neutrons lose their energy in small steps and slowly reach the capture resonance energy of an element for which the transmutation rate is maximum. Large amounts of 99Technetium, 129Iodine, or other long-lived fission products were destroyed on the periphery of the core where the neutrons have the right energy. It was suggested that the process could also be used for the production of radioisotopes for medical applications. In conclusion, nuclear transmutation can ease the waste-management problem if adequate partition (chemical separation of isotopes) can be achieved and highly efficient techniques can be developed on an industrial scale. Nevertheless, the need for geological repositories of radioactive waste will not go away completely, as in practice, neither partitioning nor transmutation can be implemented with the ideal 100% efficiency [56].

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[33] Lovley DR, Phillips EJP. Reduction of uranium by Desulfovibrio desulfuricans. Appl Environ Microbiol. 1992;58:850–6. [34] Prakash D, Gabani P, Chandel AK, Ronen Z, Singh OV. Bioremediation: a genuine technology to remediate radionuclides from the environment. Microb Biotechnol. 2013;6:349–60. doi: 10.1111/1751-7915.12059. [35] Lovley DR, Phillips EJ. Reduction of chromate by Desulfovibrio vulgaris and its c(3) cytochrome. Appl Environ Microbiol. 1994;60:726–8. [36] Lloyd JR. Microbial reduction of metals and radionuclides. FEMS Microbiol Rev. 2003;27:411–25. [37] Lloyd JR, Macaskie LE. A novel phosphorImager based technique for monitoring the microbial reduction of technetium. Appl Environ Microbiol. 1996;62:578–82. [38] Lloyd JR, Sole VA, Van Praagh CV, Lovley DR. Direct and Fe(II)-mediated reduction of technetium by Fe(III)-reducing bacteria. Appl Environ Microbiol. 2000;66:3743–9. [39] Wildung RE, Gorby YA, Krupka KM, et al. Effect of electron donor and solution chemistry on products of dissimilatory reduction of technetium by Shewanella putrefaciens. Appl Environ Microbiol. 2000;66:2451–60. [40] Lloyd JR, Macaskie LE. Bioremediation of radionuclide-containing waste waters. In: Lovley DR, editor. Environmental microbe-metal interactions. Washington, DC: ASM Press; 2000. p. 277–327. [41] Tamponnet C, Declerck S. Radionuclide (RN) pollution is a worldwide problem that arises from human activities. J Environ Radioact. 2008;99:773–4. [42] Kim SJ, Koh DC, Park SJ, et al. Molecular analysis of spatial variation of iron-reducing bacteria in riverine alluvial aquifers of the Mankyeong river. J Microbiol. 2012;50:207–17. [43] Rusin PA, Quintana L, Brainard JR, et al. Solubilization of plutonium hydrous oxide by ironreducing bacteria. Environ Sci Technol. 1994;28:1686–90. [44] Fendrihan S, Legat A, Gruber C, et al. Extremely halophilic archaea and the issue of long term microbial survival. Rev Environ Sci Biotechnol. 2006;5:203–18. [45] Schubert BA, Lowenstein TK, Timofeeff MN, Parker MA. Halophilic archaea cultured from ancient halite, Death Valley, California. Environ Microbiol. 2010;12:440–54. [46] Gramain A, Chong Díaz GC, Demergasso C, Lowenstein TK, McGenity TJ. Archaeal diversity along a subterranean salt core from the Salar Grande. Environ Microbiol. 2011;13:2105–21. [47] Ams DA, Swanson JS, Szymanowski JES, Fein JB, Richmann M, Reed DT. The effect of high ionic strength on neptunium (V) adsorption to a halophilic bacterium. Geochim Cosmochim Acta. 2013;110:45–57. doi: 10.1016/j.gca.2013.01.024. [48] Bader M, Müller K, Foerstendorf H, et al. Multistage bioassociation of uranium onto an extremely halophilic archaeon revealed by a unique combination of spectroscopic and microscopic technique. J Hazard Mater. 2017;327:225–32. [49] Gruber C, Legat A, Pfaffenhuemer M, et al. Halobacterium noricense sp. nov., an archaeal isolate from a bore core of an alpine Permian salt deposit, classification of Halobacterium sp. NRC-1 as a strain of H. salinarum and emended description of H. salinarum. Extremophiles. 2004;8:431–9. [50] Bader M, Rossberg A, Steudtner R, et al. Impact of haloarchaea on speciation of uranium – a multispectroscopic approach. Environ Sci Technol. 2018;52:12895–904. doi: 10.1021/acs. est.8b02667. [51] Bader M, Müller K, Foerstendorf H, et al. Comparative analysis of uranium bioassociation with halophilic bacteria and archaea. PLoS One. 2018;13(1):e0190953. doi: 10.1371/journal. pone.0190953. [52] Swanson JS, Norden DM, Khaing HM, Reed DT. Degradation of organic complexing agents by halophilic microorganisms in brines. Geomicrobiol J. 2013;30:189–98. doi: 10.1080/01490451.2012.659332. [53] Stan-Lotter H, McGenity TJ, Legat A, et al. Very similar strains of Halococcus salifodinae are found in geographically separated Permo-Triassic salt deposits. Microbiology. 1999;145:3565–74.

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[54] Vreeland RH, Jones J, Monson A, et al. Isolation of live cretaceous (121–112 million years old) halophilic archaea from primary salt crystals. Geomicrobiol J. 2007;24:275–82. [55] Nuclear Energy Agency. Transmutation of radioactive waste. 2012. Available from: http://www. oecd-nea.org/trw/. Accessed February 1, 2020. [56] Partitioning and transmutation: towards an easing of the nuclear waste management problem. Euratom EU Luxembourg, Office for Official Publications of the European Communities. European Commission; Luxembourg 2001. ISBN 92-894-1074-4.

Jeannette Marrero, Orquidea Coto and Axel Schippers

11 Metal bioleaching: fundamentals and geobiotechnical application of aerobic and anaerobic acidophiles Abstract: Bioleaching of sulfide ores has been extensively studied for more than 50  years and is applied worldwide at industrial level. However, the processing of oxide ores such as laterites lags behind in theory and application. This book chapter provides an update of the biotechnical applications of acidophiles in biomining of sulfide ores as well as will focus on the bioleaching of iron oxide minerals for the recovery of nickel, cobalt, manganese, and copper mainly. Moreover, this chapter gives an update of the main adaptation mechanisms to low pH developed by acidophiles. In addition, an overview is given about fundamentals of the mechanisms of bioleaching of sulfide and laterite ores, including aerobic and anaerobic bioprocesses, as well as examples of commercial applications and future directions for biomining with acidophiles.

11.1 Introduction Natural copper leaching processes were documented already in ancient times during the East Han Dynasty (206 BC–220 AD) and Song Dynasty (960–1271 AD) [1]. However, it was not until 1947 that these phenomena were attributed to bacteria, through the discovery of the iron-oxidizing proteobacterium Acidithiobacillus ferrooxidans by the pioneer work of Colmer and Hinkle [2]. This was the first characterized ferrous ironoxidizing acidophile, which was firstly named Ferrobacillus sulfooxidans and lateron Ferrobacillus ferrooxidans and Thiobacillus ferrooxidans. Historically, Acidithiobacillus ferrooxidans, together with the sulfur-oxidizing proteobacterium Acidithiobacillus thiooxidans, which was the first described acidophile [3], has been the most studied microorganism in biohydrometallurgy and the related environmental field. Biomining or biohydrometallurgy are branches of geobiotechnology dedicated to processing of ores, metal-containing materials, and solutions by means of microorganisms to recover metals. Biomining is a general term, covering both the main geobiotechniques, named as bioleaching and biooxidation. These two bioprocesses are alternatives to conventional mining and metallurgical industrial processes, which discharge large amounts of carbon dioxide, are capital intensive, and require energyconsuming steps as well as in many cases with harsh impact on the environment. Bioleaching is the extraction of metals by means of microorganisms. Copper, nickel, zinc, and uranium are the main metals recovered by bioleaching. Leaching

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microorganisms are also applied to dissolve interfering metal sulfides (MSs) in gold and silver-bearing refractory ores prior to cyanidation treatment. This process is named biooxidation because the bioleached solubilized metals are not intended to be recovered. Nowadays, the gradual depletion of high-grade metal-containing sulfide ores, the increase in the processing costs and more tighten environmental regulations have motivated the mining industry to take advantage of these microbial-driven processes to lower production costs and to aid in the development of more environmentally sound processes. In fact, the recent development in this field enables bioprocessing to compete successfully with other hydrometallurgical procedures. Additionally, biomining is increasingly recognized sometimes as the unique alternative to recover metals from low grade ores, which cannot be otherwise extracted by pyrometallurgy or hydrometallurgical methods (e.g. high amounts of toxic elements not accepted by smelters). Kennecott Utah Copper was the first to begin commercial application of bioleaching of copper sulfides in the late 1950s at the Bingham Canyon Mine near Salt Lake City, Utah, and later at the Chino mine in New Mexico, USA. Currently, biological processing is widely practiced at commercial scale for the extraction of base metals, such as copper, and nickel and zinc in lesser extents and precious metals, mostly gold, from ores and mineral concentrates. Bioleaching of sulfide minerals has been extensively reviewed and already applied at commercial scale for more than 50 years until now [4–7]. However, the processing of oxide ores, such as laterites, lags behind theory and application. In this chapter, the current progress in fundamentals and application of metal bioleaching/ biomining is reviewed with particular emphasis on bioleaching of oxide ores.

11.2 Acidophiles: habitats and adaptation to low pH Generally, microorganisms are classified as acidophiles and alkaliphiles on the basis of their optimum pH for growth [8]. Acidophilic microorganisms are present in all three domains of life and are defined as having an optimum growth pH of