Biomaterials: Biological Production of Fuels and Chemicals 9783110342420, 9783110342307

Citation preview

Luque, Xu (Eds.) Biomaterials

Also of Interest Biorefineries. An Introduction Aresta, Dibenedetto, Dumeignil (Eds.), 2015 ISBN 978-3-11-033153-0, e-ISBN 978-3-11-033153-0

Biomimetics. A Molecular Perspective Jelinek, 2013 ISBN 978-3-11-028117-0, e-ISBN 978-3-11-028119-4

Lignocellulose Chemistry Rosenau (Ed.), 2016 ISBN 978-3-11-035583-3, e-ISBN 978-3-11-035683-0

Nanocarbon-Inorganic Hybrids. Next Generation Composites for Sustainable Energy Applications Eder, Schlögl (Eds.), 2014 ISBN 978-3-11-026971-0, e-ISBN 978-3-11-026986-4

Nanocellulose. From Nature to High Performance Tailored Materials Dufresne, 2012 ISBN 978-3-11-025456-3, e-ISBN 978-3-11-025460-0

Biomaterials

| Biological Production of Fuels and Chemicals Edited by Rafael Luque and Chun-Ping Xu

Editors Prof. Rafael Luque Departamento de Química Orgánica Universidad de Córdoba Campus de Rabanales Edificio Marie Curie (C-3) Ctra Nnal IV, Km 396 Córdoba (Spain) E-14014 [email protected] Prof. Dr. Chun-Ping Xu College of Food and Biological Engineering Zhengzhou University of Light Industry Kexue road 166 Zhengzhou High-tech Zone Zhengzhou, Henan, 453000 P.R.China [email protected]

ISBN 978-3-11-034230-7 e-ISBN (PDF) 978-3-11-034242-0 e-ISBN (EPUB) 978-3-11-038358-4 Set-ISBN 978-3-11-034243-7

Library of Congress Cataloging-in-Publication Data A CIP catalog record for this book has been applied for at the Library of Congress. Bibliographic information published by the Deutsche Nationalbibliothek The Deutsche Nationalbibliothek lists this publication in the Deutsche Nationalbibliografie; detailed bibliographic data are available on the Internet at http://dnb.dnb.de. © 2016 Walter de Gruyter GmbH, Berlin/Boston Cover image: Johny87/iStock/Thinkstock Typesetting: PTP-Berlin, Protago-TEX-Production GmbH, Berlin Printing and binding: Hubert & Co. GmbH & Co. KG, Göttingen ♾ Printed on acid-free paper Printed in Germany www.degruyter.com

Contents List of contributing authors | IX Gareth M. Forde, Thomas J. Rainey, Robert Speight, Warren Batchelor, and Leonard K. Pattenden 1 Matching the biomass to the bioproduct | 1 1.1 Introduction | 1 1.2 Upstream bioprocesses | 4 1.2.1 Open systems | 4 1.2.2 Closed systems | 5 1.3 Downstream bioprocesses | 8 1.3.1 Overview of downstream bioprocesses | 8 1.3.2 Downstream bioprocessing unit operations | 9 1.3.3 Specific processing considerations for biomolecules | 10 1.3.4 Chromatography | 11 1.3.5 Stabilization and formulation | 13 1.3.6 Mechanical considerations: Pumps, valves, piping, mixing | 14 1.4 Sample bioprocesses | 16 1.4.1 Bioenergy | 16 1.4.2 Nutraceutical example: Chondroitin | 23 1.4.3 Fermentation example: Fermentation products from ligno-cellulose | 25 1.4.4 Biopharmaceutical example: Monoclonal antibodies | 31 1.4.5 Novel material example: Cellulose nanofibers | 35 1.5 Conclusion | 38 Meng Liang, Xiaowei Zhou, and Chunping Xu 2 Systems biology in biofuel | 45 2.1 Introduction | 45 2.2 The importance of systems biology | 46 2.2.1 Genomics | 46 2.2.2 Transcriptomics | 47 2.2.3 Proteomics | 48 2.2.4 Metabolomics | 48 2.2.5 Fluxomics | 49 2.2.6 Computational Methods | 49 2.3 Applicability of systems biology in biofuels | 50 2.3.1 Biodiesels | 50 2.3.2 Jet fuels | 51 2.3.3 Biobutanol | 52 2.4 Conclusions and outlook | 53

VI | Contents

Nathalie Berezina 3 Production and application of chitin | 61 3.1 Introduction | 61 3.2 Historical outline | 62 3.3 Sources | 63 3.4 Extraction and purification | 64 3.4.1 Chemical extraction | 65 3.4.2 Biological extraction | 65 3.5 Applications | 66 3.5.1 Biomedical applications | 66 3.5.2 Agricultural applications | 67 3.5.3 Materials applications | 67 3.5.4 Water purification | 68 3.6 Outlook | 68 Hui Li, Hu Zhu, Shiwei Sun, Zhimei Feng, Yajie Sun, and Wanlong Zhou 4 Biological production of welan gum | 73 4.1 Sphingans: Occurrence and structure | 73 4.2 Welan gum: structure and properties | 75 4.3 Production of welan gum | 76 4.3.1 Producing strains | 76 4.3.2 Producing conditions | 77 4.3.3 Recovery and purification of welan gum | 80 4.4 Biosynthetic pathway of welan gum | 81 4.4.1 Synthesis of the nucleotide-sugar precursors | 84 4.4.2 Assembly of the tetrasaccharide repeating unit | 85 4.4.3 Polymerization and export | 86 4.4.4 Regulation of welan gum biosynthesis | 87 4.4.5 Enzymes in other process | 87 4.5 Engineering approaches for improvement of sphingan production | 88 4.6 Applications of welan gum | 89 4.6.1 Cement systems | 89 4.6.2 Oil industry | 90 4.6.3 Other potential applications | 91 4.7 Future perspectives | 91 Han Wei, Tang Junhong, and Li Yongfeng 5 Utilization of food waste for fermentative hydrogen production | 95 5.1 Introduction | 95 5.2 Metabolic pathway of fermentative hydrogen production | 96

Contents

5.2.1 5.2.2 5.3 5.3.1 5.3.2 5.3.3 5.4 5.4.1 5.4.2 5.4.3 5.5 5.6

| VII

Process yield and conversion efficiency | 96 Metabolic pathway for fermentative hydrogen production | 97 Biohydrogen production from food waste | 99 Carbohydrate | 100 Fats | 100 Protein | 100 Pretreatment of food waste for fermentative hydrogen production | 100 Physical pretreatment | 101 Chemical pretreatment | 101 Enzymatic pretreatment | 101 Performance of biohydrogen production from food waste | 102 Prospects and challenges of fermentative hydrogen production from food waste | 103

Chao Chen and Tao Li 6 Bacterial dye-decolorizing peroxidases | 107 6.1 Introduction | 107 6.2 Biochemical properties | 109 6.3 Physiological roles of bacterial DyPs | 111 6.4 Catalytic mechanism of bacterial DyPs | 113 6.5 Structure-function relationship in bacterial DyPs | 117 6.6 Biotechnological opportunities | 122 6.6.1 Lignin valorization and fine chemicals | 122 6.6.2 Dye decolorization in wastewater treatment | 123 6.6.3 Other potential industrial applications | 124 6.7 Conclusions and perspectives | 125 Pei-Ching Chang, Hsi-Yen Hsu, and Guang-Way Jang 7 Biological routes to itaconic and succinic acids | 131 7.1 Introduction | 131 7.1.1 Succinic acid | 131 7.1.2 Itaconic acid | 133 7.2 Synthesis of succinic acid and derivatives | 134 7.2.1 Chemical synthesis of succinic acid | 135 7.2.2 Biological routes to succinic acid | 135 7.2.3 Catalytic conversion of succinic acid | 142 7.3 Synthesis of itaconic acid and derivatives | 145 7.3.1 Biological routes to itaconic acid | 145 7.3.2 Catalytic conversion of itaconic acid | 148 7.4 Applications of succinic acid and derivatives | 148

VIII | Contents

7.5 7.6

Applications of itaconic acid and derivatives | 150 Conclusions | 153

Lei Chen, Xingxun Liu, and Ka-Hing Wong 8 Novel nanoparticle materials for drug/food delivery-polysaccharides | 159 8.1 Introduction | 159 8.2 Nanoparticles for delivery systems | 160 8.3 Polysaccharides and their nanoparticles | 161 8.3.1 Nonpolyelectrolyte polysaccharides | 163 8.3.2 Positively charged polyelectrolyte polysaccharides | 167 8.3.3 Negatively charged polyelectrolyte polysaccharides | 168 8.3.4 Hyperbranched polysaccharides | 170 8.3.5 Other polysaccharides | 171 8.4 Nanoparticle preparation based on polysaccharides | 171 8.4.1 Covalent crosslinking polysaccharide nanoparticles | 171 8.4.2 Ionic crosslinking polysaccharide nanoparticles | 172 8.4.3 Polyelectrolyte complexing polysaccharide nanoparticles | 173 8.4.4 Self-assembly polysaccharide nanoparticles | 173 8.5 Applications of polysaccharide-based nanoparticles | 174 8.5.1 Medical applications | 174 8.5.2 Food applications | 176 8.6 Conclusions | 177 Index | 191

List of contributing authors Warren Batchelor Bioresource Processing Institute of Australia (BioPRIA) Department of Chemical Engineering Monash University www.monash.edu Chapter 1 Nathalie Berezina Ynsect 1 rue Pierre Fontaine 91058 Evry, France [email protected] Chapter 3 Pei-Ching Chang Material and Chemical Research Laboratories Industrial Technology Research Institute 321 Kuang Fu Road, Hsinchu, 30011, Taiwan Chapter 7 Chao Chen School of Pharmacy University of Connecticut Storrs, CT 06269, United States Chapter 6 Lei Chen The Key Laboratory of Industrial Biotechnology Ministry of Education School of Biotechnology Jiangnan University Wuxi 214122 China Chapter 8 Zhimei Feng Center for Bioengineering and Biotechnology China University of Petroleum (East China) No 66, Changjiang West Road Qingdao Economic & Technical Development Zone Qingdao, Shandong China, 266580 Chapter 4

Gareth M. Forde Energy and Process Engineering Discipline School of Chemistry, Physics and Mechanical Engineering Science & Engineering Faculty Queensland University of Technology [email protected] and All Energy Pty Ltd. www.allenergypl.com.au Chapter 1 Hsi-Yen Hsu Material and Chemical Research Laboratories Industrial Technology Research Institute 321 Kuang Fu Road, Hsinchu, 30011, Taiwan Chapter 7 Guang-Way Jang Material and Chemical Research Laboratories Industrial Technology Research Institute 321 Kuang Fu Road, Hsinchu, 30011, Taiwan [email protected] Chapter 7 Tang Junhong College of Materials and Environmental Engineering Hangzhou Dianzi University Hangzhou 310018 China Chapter 5 Hui Li Center for Bioengineering and Biotechnology China University of Petroleum (East China) No 66, Changjiang West Road Qingdao Economic & Technical Development Zone Qingdao, Shandong China, 266580 Chapter 4

X | List of contributing authors

Tao Li Sustainable Technology Division, NRMRL Office of Research and Development US EPA, Cincinnati, Ohio, USA, 45268 Chapter 6 Meng Liang School of Food Science and Biotechnology ZhengZhou University of Light Industry ZhengZhou Henan, China Chapter 2 Xingxun Liu Institute of Food Science and Technology (IFST) Chinese Academy of Agricultural Science (CAAS) Beijing China Chapter 8 Leonard K. Pattenden Biosciences and Biotechnology School of Applied Sciences RMIT University Chapter 1 Thomas J. Rainey Energy and Process Engineering Discipline School of Chemistry, Physics and Mechanical Engineering Science & Engineering Faculty Queensland University of Technology [email protected] Chapter 1 Robert Speight Energy and Process Engineering Discipline School of Chemistry, Physics and Mechanical Engineering Science & Engineering Faculty Queensland University of Technology Chapter 1

Shiwei Sun Center for Bioengineering and Biotechnology China University of Petroleum (East China) No 66, Changjiang West Road Qingdao Economic & Technical Development Zone Qingdao, Shandong China, 266580 Chapter 4 Yajie Sun Center for Bioengineering and Biotechnology China University of Petroleum (East China) No 66, Changjiang West Road Qingdao Economic & Technical Development Zone Qingdao, Shandong China, 266580 Chapter 4 Han Wei College of Materials and Environmental Engineering Hangzhou Dianzi University Hangzhou 310018 China Chapter 5 Ka-Hing Wong Food Safety and Technology Research Centre Department of Applied Biology and Chemical Technology The Hong Kong Polytechnic University Hung Hom, Kowloon, Hong Kong China Chapter 8 Chunping Xu School of Food Science and Biotechnology ZhengZhou University of Light Industry ZhengZhou Henan, China Chapter 2

List of contributing authors

| XI

Li Yongfeng School of Forestry Northeast Forestry University Harbin 150060 China Chapter 5

Xiaowei Zhou School of Food Science and Biotechnology ZhengZhou University of Light Industry ZhengZhou Henan, China Chapter 2

Wanlong Zhou Center for Bioengineering and Biotechnology China University of Petroleum (East China) No 66, Changjiang West Road Qingdao Economic & Technical Development Zone Qingdao, Shandong China, 266580 Chapter 4

Hu Zhu Center for Bioengineering and Biotechnology China University of Petroleum (East China) No 66, Changjiang West Road Qingdao Economic & Technical Development Zone Qingdao, Shandong China, 266580 Chapter 4

Gareth M. Forde, Thomas J. Rainey, Robert Speight, Warren Batchelor, and Leonard K. Pattenden

1 Matching the biomass to the bioproduct Summary of up- and downstream bioprocesses 1.1 Introduction As noted by the science journalist Robin Williams, “The 20th century was the century of physics, the 21st century will be the century of biology”. Knowledge of biological systems started firstly with observation or documentation, followed by understanding, then finally utilization or biomimicking. Whilst humans have been employing bioprocesses since as early as 7000 BC (as evidenced by Neolithic fermentation jars), our detailed understanding of biological processes in terms of genomics, metabolomics, and proteomics is very recent. For a production process to be economically viable, the availability of the feedstock, the unit operations and the product must be understood. Lower value products (e.g. livestock feed), bulk commodities (e.g. sugar) or products with a number of competing sources (e.g. electricity) require scale, low cost feedstocks (e.g. sourced from broad-acre, agri-waste or animal tissues), and either low cost or efficient unit operations (e.g. high yields, low utility requirements). Higher value biomolecules (e.g. enzymes; biopharmaceuticals) can bear higher cost feedstocks (e.g. from fermentation) with higher cost intensity unit operations (e.g. centrifugation, spray drying and chromatography). Fig. 1.1 shows some examples of biomass feedstocks or products and their associated value as a function of cellulose and hemicellulose content. Their abundance in many forms of biomass and their attractiveness for fuel drive down their price, but there is substantial opportunity for further value adding. Biomass is inherent and, in the vast majority of cases, nontoxic as biological processes create products which serve as feedstocks for other processes (with notable exceptions, such as venoms and pathogens, representing extremely low percentages of the total global biomass). Hence, a well-engineered bioprocess should be “benign by design”. Bioprocess engineering is defined as the design and development of processes for the manufacture of products from biomass or via biological processes. By using the inspiration offered by biological systems, bioprocess engineering should strive for creating circular processes where everything other than the end product can be a feedstock for another process. The inspiration to address many of society’s current challenges using biology has spawned the area of biomimetics, where an engineered biological system aims to mimic the inherent advantages of natural biological systems. An example is where humans look to biology for examples of biofixation of carbon dioxide (CO2 ). CO2 is taken from a gaseous phase and converted into

2 | 1 Matching the biomass to the bioproduct

Hemi- and cellulosic content

70% 60%

Recycled waste wood

Clean chipped Bagasse wood Livestock feed

Functional Foods

Nutraceuticals Foods

50% 40% 30%

Thermal fuel

Commodities and animal Bulk raw grain Algae (cultured)

20% Bulk raw oil seeds

10%

–$100

$100

$1000

Food

Gold $10k

Protein

$100k

Chondroitin sulphate

$1 mil

Amino acids

$10 mil

Value $/t Fig. 1.1: Correlation of dry weight cellulose and hemicellulose content to value in $ per ton. Note that the x-axis is logarithmic.

biomass and biological compounds (e.g. carbohydrates, lipids, DNA, protein), then this biomass is reused as feedstock for products such as transport fuel, energy, polymers, food, food additives and fine chemicals. Whilst such systems may have inherent inefficiencies, if the net environmental impact is zero, then the economics may be improved through scale and smart engineering. Industry must find ways to achieve deep decarbonization of how energy, food and products (polymers, drugs, etc.) are created, that is, to stop the reliance on fossil fuels. Biomass processing coupled to innovative downstream processing provides one significant opportunity to decarbonize human society. Use of life cycle analyses (LCAs) can provide insights for making long term decisions on sustainable processes. For example, the biopharmaceutical industry has one of the highest waste to product ratios of any industry (kg waste generated per kg product produced) compared to the oil and gas industry which has the lowest waste to product ratio. This high waste ratio is caused by the extensive use of disposables and cleaning requirements warranted in the manufacture of a biopharmaceutical. One option to improve the sustainability of bioindustries (refer Fig. 1.2) is anaerobic digestion to produce bioenergy from organic byproducts, rather than chemical treatment or off-site disposal. Other examples include the recycling of nutrients or isolation of fractions from waste streams for other purposes such as carbon-fiber production from waste lignin. This chapter will consider which biomass feedstocks are most suited for the manufacture of bioproducts (e.g. bioenergy, chemicals, vaccines) via appropriate bioprocess engineering unit operations. For example, a low value biomass (chipped wood)

1.1 Introduction

| 3

requires simple, low cost and large scale unit operations whilst the manufacture of a high value bioproduct (e.g. a biopharmaceutical or functional food) utilizes a larger number of smaller scale and high cost unit operations to achieve appropriate purities. The chapter will consider upstream processes (the creation of the biomass), downstream processes (the separation of the target biomolecules or value adding stages (refer Fig. 1.2 for a sample flow diagram) and finally present some examples of the creation of products from biomass.

Raw materials

Cells

Media preparation /sterilisation

Inoculum

Bioreaction/fermenters/ cell growth

By-products (e.g. to anaerobic digestion to generate bio-energy)

Separation of cells from supernatant (harvest)

Liberation (lysis/cell disruption)

Product concentration (e.g. filtration) and purification (e.g. chromatography)

Final product formulation, packaging, qa/qc

Fig. 1.2: Block diagram of a bioprocess for the manufacture of a purified biomolecule via fermentation. The stage of product concentration and purification represents approximately 50 to 80 % of total processing costs.

4 | 1 Matching the biomass to the bioproduct

1.2 Upstream bioprocesses Upstream biomass generation can be in “open” or “closed” systems, where open systems are traditionally utilized for the generation of food and energy crops using agricultural or cropping processes, whilst closed systems are routinely characterized by monocultures (e.g. bacterial, yeast) achieved via the use of sterile procedures with defined or semi-defined media.

1.2.1 Open systems Food, forestry and energy crops are typically based around sugars, starch and lignocellulosic material. Harvesting operations for many food industries involve leaving some nonfood material in the field (e.g. stubble from grain or leaves from sugarcane) while the harvested material is separated into (i) food and (ii) nonfood material. The greater the chemical complexity of the nonfood material (e.g. lignocellulosics), the more challenging is the processing and value adding. Similarly timber is harvested, sawn to produce lumber and the residue is chemically complex which results in much of it being burned (Section 1.4.1). Notwithstanding, open systems can produce high value materials – an example is provided in Section 1.4.5. Typical sources of biomass from open systems: – Food agriculture – Sugarcane industry – Grain industry – Fruit and vegetable – Other agriculture – Cotton – Dedicated energy crops (e.g. miscanthus) – Forestry – Timber – Bark – Woodchip – Leaves, branches, stumps The products from biomass are wide (in order of increasing value): – Electricity – Building products – Transport fuels – Food

1.2 Upstream bioprocesses

–

–

|

5

Pulp and paper products – Newsprint – Photocopier paper – Tissue Niche materials (e.g. microfibrillated cellulose)

Typical upstream operations include: – Planting – Crop maintenance (e.g. fertilizing, spraying, pest control) – Harvesting – Transportation – Coarse separation of the plant into different components (e.g. grain from husk and bark from the timber) by mechanical processing – Combusting part of the feedstock may occur to assist upstream processing (e.g. sugarcane)

1.2.2 Closed systems For closed systems biomass is composed of cells which may, from an engineering perspective, be considered as a highly organized molecular factory, see Fig. 1.3. Our modern understanding of cells is not perfect clones of the same cell, but rather colonies of interacting cells with specialized functions.

Environment:

Temp pH Dissolved oxygen Shearing forces– mixing, convection Heat and mass transfer

Vector Cell Medium Macronutrients (C, N, H, O, S, P, K, Mg) Micronutrients

Redox potential (Oxidative stress– apoptosis pathways)

Fig. 1.3: Upstream or BIOREACTION section: Selection and conversion.

6 | 1 Matching the biomass to the bioproduct

Deloitte Touche Tohmatsu [1] reports that global fermentation industries are valued at over US $ 127 billion per annum made up of ethanol (87 % of value) with the next two largest groups being amino acids (8.7 %) and organic acids (2.8 %). The projected annual growth to 2020 (excluding alcohols) for fermentation products is 6.5 %, in which the highest growth area is polymers. The primary characteristics of a cell are the cell membrane (or plasma membrane), cytoplasm and organelles and the nuclear region. The similarities between all cells are: 1. Cell membrane 2. Contains DNA 3. Composed of the same basis chemicals: carbohydrates, proteins, nucleic acids, minerals, fats and vitamins 4. All cells regulate the flow of nutrients and wastes that enter and leave the cell 5. All cells reproduce and are the result of reproduction 6. All cells require a supply of energy 7. All cells responding to stimuli are highly regulated by elaborate sensing systems Biomolecules come from a vast number of sources: – From all types of cells imaginable – Synthesized (peptides and oligos) Biomolecules have an even larger number of applications, in increasing value: – Livestock feed – Human food and beverage – Function foods e.g. enhanced foods – Nutraceuticals e.g. amino acids, proteins – Fine chemicals – Enzymes – Biopharmaceuticals e.g. vaccines, monoclonal antibodies, gene therapy agents Typical upstream unit operations include: – Medium formulation – Sterilization – Cell culturing – Propagation – Biomass production – Metabolite biosynthesis – Aggregation/harvesting – Hydrolysis – Biotransformations

1.2 Upstream bioprocesses

| 7

One of the main actions for bulk biomass and cell cultures is the reduction of water content. The composition of a typical cell is 70 % or more water with the other mass (dry weight) consisting of the molecules outlined below (Tab. 1.1). Tab. 1.1: Moisture contents of biomolecules. Class

% dry weight

Constituents

Protein Carbohydrates Lipid RNA DNA Small molecules

55 10 9 20 3 3

amino acids sugars fatty acids, glycerol, phosphate purines, pyrimidines, phosphate, and ribose purines, pyrimidines, phosphate, and deoxyribose metabolic intermediates and inorganic ions (e.g. metals ~1 %)

Big, glycosylated, complex molecules are more likely to be sourced from biomass, whilst smaller, simpler molecules are likely to be generated by synthetic means (Tab. 1.2). Tab. 1.2: KPIs of cellular systems for rDNA protein production. Key Performance lndicator High µ Expression Ieveis Low-cost media Protein folding Simple posttranslational processing Complex posttranslational processing Low proteolyic degradation Secretion Safety Sealability Posttranslational processing required (e.g. glycosylation) Posttranslational processing and secretion NOT required (e.g. glycosylation)

E. coli

Yeast

lnsect

Plant

Mammalian

4 4 4 1 0 0 1.5 0 3 4

3 3 4 1.5 2 0 2 3 4 3.5 26

0.5 1.5 0 3.5 3 0 3 3 4 3 21.5

0.5 1.5 1 4 4 2 3 4 3 2 25

0.5 1.5 0 4 4 4 3 4 2 2.5 25.5

21

15.5

15

13.5

21.5

8 | 1 Matching the biomass to the bioproduct

1.3 Downstream bioprocesses 1.3.1 Overview of downstream bioprocesses Downstream processing refers predominantly to the physical and chemical processing to extract, purify then concentrate the target molecule from the water and biomass created in the upstream stage. The processing of the bulk feed material (e.g. whole cells suspended in cell culture) must ensure that an appropriate chemical structure and biochemical activity is maintained whilst ensuring the entire process is economically viable. The process must have sufficient yield and appropriate capital and operating costs. The level of processing depends on the difficult degree of purification, the intended use of the final product and the final required specification. Industrial fuels, chemicals and materials will generally require fewer processing steps than a product that will be consumed by humans or animals. Even within human use products, the level of bioprocessing increases as the therapeutic use of the molecule increases (i.e. administered intravenously rather than orally and must have a specific efficacy rather than general effects). For example, some food supplements may be simply dried and milled, whilst a biopharmaceutical may undergo a dozen or more downstream unit operations to achieve the required purity and sterility. The principal aims of downstream processing are to: – maximize target product/biomolecule yield – minimize the number of unit operations (because product loss occurs at each purification step) – minimize processing time and cost – maintain biomolecule activity/integrity by preventing degradation (due to temperature, pH, chemical reactions, enzyme hydrolysis, shear, UV degradation, etc.) – ensure product quality in terms of specified product composition and reproducibility (i.e. no variation between doses, batches or during storage) – provide a product with an acceptable concentration, volume, and guaranteed activity – provide a safe product that is, for example, free of toxins (endotoxins/lipopolysaccharides), viruses, chemical contaminants and pathogens – where required, uses a validated process such as a current Good Manufacturing Process (cGMP) The aims of downstream processing can be counterproductive, with a compromise or economic optimization being required. For example, whilst chromatography may provide a fast and single unit operation for the purification of a recombinant protein, it may result in a cost impost which is unacceptable (due to equipment, resin, buffer media, and operating costs). As a further example, if the aim is to create a high purity and high biological activity enzyme, more unit operations contribute to achieving a

1.3 Downstream bioprocesses |

9

Monoclonal antibodies (Insulin) Neutraceuticals

Vaccines

Penicillin Enzymes amino acids Food & beverage Energy 100

DNA/RNA

Therapeutic proteins

(Cancer prevention/treatment & immunologic diseases)

Gold

Difficulty to produce

higher purity but often reduce biological activity through increased opportunity for degradation and denaturation. Downstream processing is routinely utilized for the manufacture of food supplements, nutraceuticals and biopharmaceuticals. Biopharmaceuticals represent a new generation of therapeutics of rapidly increasing importance: in 2002 the total pharma market was US $ 390 billion, of which biopharmaceuticals accounted for 7 % (US $ 27.3 billion); in 2005 it was 12 % (US $ 70.8 billion); in 2010 about 50 % of drugs in development were biopharmaceuticals [2]; in 2013 it was estimated to represent around 20 % (approximately US $ 199.7 billion) with the predicted biopharma market in 2020 as high as US $ 490 billion [3]. The global nutraceuticals product market was US $ 142.1 billion in 2011 and is expected to reach US $ 204.8 billion by 2017, with ingredients for the nutraceutical industry valued at US $ 33.6 billion [4]. The functional food and beverage market reached US $ 93 billion in 2011. Fig. 1.4 shows how the value of a biomolecule increases via additional processing and as the therapeutic requirements increase in complexity. That is, small batch size therapeutic proteins for low frequency disease states and gene therapy molecules are much more expensive to make than large production runs of vaccines and monoclonal antibodies.

104

Selling price $/kg

1010

Fig. 1.4: Value in US $ per kg (x-axis) versus indicative difficulty to produce different products sourced from biomass and biomass-derived sugars. As the requirement for high purity increases, so too does the number of unit operations required to produce the final product. Note that the x-axis is a logarithmic scale.

1.3.2 Downstream bioprocessing unit operations Listed below are the most common unit operation employed in downstream processing. This is not an exhaustive list, nor does it detail the specifics of each unit operation as this is left for other text books dedicated to these topics. The list provides an indicative ranking of unit operations from cheapest to most expensive on a $/ton processed basis. The first few unit operations (coarse size separation, comminution and biomass

10 | 1 Matching the biomass to the bioproduct

dewatering) are the only unit operations routinely used for biomass energy applications as further processing is usually too expensive. To make a food supplement or high purity biomolecule injectable a large number of these unit operations may be required. Typical downstream¹ unit operations include: 1. Coarse size separation: sieving, sorting, sizing decks 2. Comminution: crushing, grinding, milling 3. Biomass dewatering: centrifugation, gravity separation, flocculation, rotating drum 4. Cell lysis/disruption or liberation: homogenization/pressure, chemical cell wall rupture 5. Extra-cellular product concentration: (a) Mechanical: tangential flow filtration, centrifugation (b) Evaporative: spray drying, freeze drying 6. Purification: (a) Ultrafiltration (b) Chromatography (c) Precipitation (e.g. via the use of salts) (d) Extraction (e.g. liquid-liquid) (e) Crystallization (f) Dia-filtration/dialysis (g) Sterile filtration 7. Stabilization/formulation/vialing/dosing/packaging

1.3.3 Specific processing considerations for biomolecules Improving the economics of the production of a biomolecule is not only about the upstream volumetric yield (i.e. grams of biomolecule per liter of cell culture) and upstream specific yield (i.e. grams of biomolecule per gram of biomass) but also: – time yield, for example, grams per hour of production time (bioprocessing plants are expensive to construct and operate, hence efficient utilization is paramount for a profitable biomolecule production process), – bioactivity (i.e. percentage of biomolecules that are biologically active), – cell and product substrate yield e.g. grams of cells and/or secondary metabolites produced per gram of glucose or other fermentable sugar consumed (or more generally, the yield of carbon in the product compared to the carbon source used), – cell and product macro-/micronutrient yields e.g. where the creation of cells and/or secondary metabolites depends upon the presence of a particular additive

1 The first four items are sometimes referred to as midstream.

1.3 Downstream bioprocesses

–

| 11

being in the growth media, the efficient use of this additive is vital for a process to be economically viable, and maintaining acceptable yields and bioactivities for all downstream unit operations.

As mentioned in Section 1.3.1, aims of downstream processing are sometimes counterproductive, especially from an economic perspective. For example, when making a dietary supplement processing the biomolecule to a purity that is fit for purpose may be more economically viable than achieving the highest purity possible. This point is emphasized by the example nutraceutical product chondroitin presented in Section 1.4.2, where the profit after tax for the “value adder” (stage in the supply chain where the target biomass is isolated) equates to 36 %, compared to 9 % for the manufacturer and 16 % for the retailer (refer Fig. 1.5). Generally, as purity requirements increase for a biomolecule, so too does the number of unit operations (or processing stages) employed. Additionally, certain unit operations or processes may destroy the target biomolecule, including: – Shearing e.g. pumps and mixing – Hydrolysis or the cleavage of molecular bonds e.g. via enzymes – Inhibition via oxidation or chelation with metal ions – Irreversible aggregation/precipitation – Extremes of pH Hence, bioactivity yield refers to both retaining the target molecule as well as the biological activity of the molecule (i.e. the amount of the molecule that it is present and that can perform its function). Tab. 1.3 shows the relationship between the number of stages or unit operations and the percentage bioactivity yield. Tab. 1.3: Overall bioactivity (%) for multiple stage processes. Percentage (%) bioactivity yield at each stage

After 3 stages

After 5 stages

After 10 stages

99 95 90 80

97 86 73 51

95 77 59 33

90 60 35 11

1.3.4 Chromatography Chromatography has been singled out as a unit operation of specific interest for the manufacture of higher value biomolecules. It is particularly important in the biopharmaceutical industry, however is not widely utilized in other process industries due

12 | 1 Matching the biomass to the bioproduct

to its batch-wise nature. Chromatography separates mixtures into components by passing a fluid (i.e. mobile or pumpable phase containing molecules in suspension) through a bed of adsorbent (i.e. stationary phase), followed by elution of the target molecule. Of particular importance within a bioprocessing context is High Pressure Liquid Chromatography (HPLC). The choice of the stationary phase and consequently the type of chromatography depends on the nature of the solutes and process goals. The column is the mechanical device which holds the adsorbent in place and is normally rated to a specific operating pressure. Separation is achieved on the basis of: – Charge (ion exchange or IE; most common in the area of protein purification) – Size – Hydrophobicity – Affinity (highly specific molecule to ligand adsorption/desorption). Chromatography uses the following mode of operation: 1. Equilibration (washing of the column with equilibrating/running buffer) 2. Sample is loaded (with associating binding to the adsorbent) 3. Washing to remove contaminants until absorbance returns to base line 4. Elution (recovering the target molecule) 5. Cleaning/regeneration The process is then started again until the adsorbent no longer provides an acceptable performance, which could be overpressure due to clogging or compression of the adsorbent, a reduction in binding capacity (g target/mL adsorbent) or lack of specificity for the target biomolecule. The Key Performance Indicators (KPIs) of chromatography are: 1. Retention factor: used to describe the migration rate of an analyte through a column. It should not be too slow so as to impact cycle time nor too fast so as to result in low binding capacity or low purity. It is also known as the capacity factor. 2. Selectivity factor: describes the degree of separation between molecules. 3. Resolution: a combination of the degree of separation between the peaks eluted from the column (selectivity), the ability of the column to produce narrow, symmetrical peaks (efficiency) and the amount (mass) of sample applied. It is defined as the distance between peak maxima compared with the average base width of the peaks. 4. Partition coefficient: concentration of analyte in the stationary phase divided by the molar concentration of the analyte in the mobile phase. It is also known as the equilibrium constant. Scale up of chromatography is achieved by firstly optimizing the purification scheme on the laboratory scale, then increasing one of the following parameters (in order of approximate preference): column diameter (cm), column volume (L), sample load (g),

1.3 Downstream bioprocesses

|

13

and finally volumetric flow rate (l/hr). The following should be endeavored to be kept constant: bed height (cm), linear flow rate (cm/h), and sample concentration (g/l). A common scale-up issue is increasing pressure drop with increasing linear flow rate leading to chromatography medium (i.e. the solid adsorbent) deformation. Hence, industrial chromatography systems are routinely less than 1.0 m in height. Liquid distribution across large bed diameters is also a key challenge for scale-up (i.e. prevention of channeling, ensuring an even binding density throughout the bed, complete washing and elution of the column).

1.3.5 Stabilization and formulation Selecting the correct stabilization and formulation for biomolecules is vital as suitable vialing, labeling, dosing and packaging is critical for storage, distribution and at the point of use. Maintenance of the structural integrity of biomolecules, and in particular therapeutics, is essential for its physiological and pharmacological efficacy. Biomolecule stabilizing strategies include: 1. Native structure stabilization (excipients/molecular engineering) 2. Prevention of aggregation (excipients) 3. Avoid or block unwanted hydrophobic surfaces (excipients and avoidance of headspace) and 4. Reduce shear forces (avoid having a headspace) The most common long term storage options for biomolecules are in solid form (e.g. freeze drying/lyophilization) or liquid form. For freeze drying, excipients are added to protect biomolecules during freezing/drying and thawing/rehydrating. For freeze drying, molecular mobility is drastically reduced thereby preventing or reducing the majority of the deactivation mechanisms: oxidation, aggregation, hydrolysis (breaking down of the chemical structure), and deamidation (nonenzymatic covalent modification which occurs to asparagine and glutamine). One negative of freeze drying is that liquid buffer needs to be added to re-suspend the dry powder before administration. Liquid solutions of biomolecules are commonly a formulation with excipients in a pH buffered. A key limitation of liquid solutions is the requirement for continuous refrigeration until the point of administration (e.g. a cold chain requiring a storage temperature of 2–8 °C; freezing generally damages biomolecules due to creation and melting of solid ice crystals). Excipients are auxiliary substances used in suspensions with examples including buffering agents, isotonicity modifiers, preservatives, stabilizers and complexing agents. Tab. 1.4 provides additional information on excipients. The use of excipients can be a delicate balance between achieving maximum shelf life, maximum bioactivity and minimum pain for the recipient (e.g. pain can result if the salt concentration is either too low or too high). Crystallization is another stabilization method but used for niche applications in the areas of formulation of proteins

14 | 1 Matching the biomass to the bioproduct

for X-ray crystallography, pharmaceuticals, and antibiotics. Crystallization operates at low temperatures to minimize thermal degradation and prevent resolubalization with associated high concentrations of the biomolecule being required. Optimization of crystallization conditions performed via empirical experimentation as phase diagrams and kinetic info are usually not available for new and novel biomolecules. Crystals are ultimately recovered via centrifugation or filtration. When choosing the vial and packaging options, consideration must be given to the light sensitivity of the molecule, diffusion of small molecules/gases and heat/cold through the packaging, usability at point of use, recyclability of the packaging, packaging weight, and packaging cost. Tab. 1.4: Examples of excipients and associated purposes. Excipient

Purpose

Example

Buffering agents

Target pH maintenance

Sodium phosphate (PBS, pH 7.4), sodium bicarbonate, sodium citrate, sodium acetate.

Isotonicity modifiers

Minimize cell damage and pain

Glycerin (16 mg/ml), sodium chloride (7 mg/ml), Target osmolality of 285 mOsmol/kg.

Preservatives

Antimicrobials

Phenol, m-cresol, methylparaben, chlorobutanol, benzyl alcohol, sorbic acid, potassium sorbate, benzoic acid, chlorocresol. Crystal growth is a main issue.

Stabilizers

Impart stability to particles or the entire suspension

Metal ions (zinc, calcium), salts, organic molecules. May be used to induce crystallization.

Complexing or formulation agents

Create stable complexes or particle formulations

Protamine sulfate (also inhibits protease activity). Wetting agents, surfactants, colloids, electrolytes, viscosity modifiers.

1.3.6 Mechanical considerations: Pumps, valves, piping, mixing Biomolecule manufacturing plants have unique requirements in terms of selecting the correct mechanical equipment and materials of construction. For example, pumps and valves are a major source of contamination in bioprocesses. Ball valves and centrifugal pumps that are static and not continuously in operation will generate “dead” areas where water, process liquid or biofilms can accumulate. The US Food and Drug Administration (FDA) notes that firms should install a drain from the low point in equipment housing.

1.3 Downstream bioprocesses

| 15

Pumps can burn out and parts can wear resulting in direct contamination (e.g. metal shavings) and failure has the potential to disrupt a process, resulting in contamination via dead zones in the process equipment. Consideration needs to be given to shear forces and pressure changes imposed during pumping of cells and bioproducts suspensions. This pressure change issue is of concern to viable cells pumped through heat exchangers external to a bioreactor. Rotary or positive displacement pumps (as opposed to centrifugal pumps) are almost exclusively used in pharmaceutical/biotech industries, utilizing electrical drives with mechanical seals or magnetic drives. Specific examples of rotary pumps include: 1. Gear pumps. These pumps are comparably continuous, produce nonpulsating flow and they are good for viscous material. 2. Lobe pumps. Uses rotating lobes to direct the flow. These pumps reduce shear effects, can drive large solids and slurry-laden media, are suitable for high viscosity liquids, have high efficiency, are corrosion resistant, and have high reliability. 3. Diaphragm pumps. These pumps are well suited for sterile/aseptic applications as the diaphragm separates the pump chambers and the pumped material. They can be air or hydraulically driven. 4. Screw pumps. Twin screw pumps are generally more reliable and result in fewer blockages (as opposed to single screw), however, metal-on-metal wearing is a source of contamination. 5. Piston pumps. Used for very accurate pumping and dispensing of a wide range of fluids. May only have one moving part hence reduces change of blockages or valve failures. Well suited to corrosive and aggressive media; suitable for high discharge pressure applications. The valve closing mechanism must be isolated from the contents of the pipe, that is the mechanical mechanisms must be isolated from the process fluid. Diaphragm valves are most commonly used in bioprocesses, such as diaphragm valves (or pinch valves) and weir-type diaphragm valves. Diaphragm valves can wear and ultimately fail during service and so routine inspection is required as well as replacing the diaphragm. Ball, butterfly and plug valves are commonly used on the utility side to reduce costs (e.g. chilled water, cooling water, hot water, steam), but are not ideal for liquids that contact the process fluid. Bioprocess facilities should have no “dead-legs”, which refers to areas that do not experience constant process flows (e.g. t-sections off the main pipe run that end in a blind flange). A design rule commonly used is to ensure that the unused portion of a pipe is not greater in length than six diameters of the unused pipe measured from the axis of the pipe in use for hot circulating streams (75–80 °C). For colder systems (65–75 °C) any dead-legs have potential for the formation of a biofilm and should be eliminated or have special sanitizing procedures. Bioprocess facilities should have no threaded fittings. All pipe joints should utilize sanitary fittings or be butt welded.

16 | 1 Matching the biomass to the bioproduct

A firm’s procedures for sanitization, as well as the actual piping, should be reviewed and evaluated during the design stages. For mixing, shear and power considerations are of primary concern. The main mechanical device selection relates to the impeller. Two common types are: – Flat bladed/Rushton impeller: exhibits higher shear, higher mass/energy transfer. Normally suited to bacteria, yeast and the preparation of suspensions and buffers. – Marine impeller: lower shear. Better suited for mammalian cell cultures. Surfaces in contact with process fluids must be constructed from easily cleaned materials of construction (e.g. stainless steel 316; appropriate polymers) and be resistant to the temperature variations and cleaning chemicals used in the process. Copper and copper alloys should not be used; nor should polymers that generate pyrogens (feverproducing agents).

1.4 Sample bioprocesses 1.4.1 Bioenergy Replacing fossil fuel based energy with renewable energy is essential to minimize global warming. Around 10 % of global energy needs are derived from bioenergy [5]. Bioenergy takes many forms however. Traditional burning of fiber (e.g. from wood) for household heating and cooking remains the largest form of bioenergy generation worldwide. Bioenergy is a major contributor to renewable electricity and transportation in many countries, such as Australia. The main bioenergy bioproducts considered here are electricity, biogas, ethanol, biodiesel and biocrude oils. This subsection elucidates the relationship between feedstocks, the bioproducts and the processes to make them. Although wind, hydroelectricity and solar power are significant sources of renewable energy, bioenergy is likely to remain a key technology for achieving greenhouse gas reduction targets. Biomass is well suited for producing liquid fuels for combustion in reciprocating engines used for transportation. Unlike many other forms of renewable energy, liquid biofuels can be blended with the existing transportation fuel supply and delivery infrastructure, allowing a gradual transition to a renewable fuel supply. It is the only renewable energy technology which has the potential to be carbon negative by absorbing carbon dioxide out of the atmosphere and returning some of it to the soil in the form of solid residue (char). First generation biofuels involve edible feedstocks and are produced by conventional processes. These feedstocks are generally sugars (e.g. sugarcane molasses), starches (corn/maize) and vegetable oils that can be readily converted into liquid fuels. These raw materials have been cultivated by mankind for thousands of years as a food source and consequently have also been explored in depth as a fuel source. To

1.4 Sample bioprocesses

|

17

this end, first generation feedstocks are grown on arable land. Using arable land to make a fuel feedstock renders it incapable of producing food, and thereby reducing food supply and potentially increasing world food prices. This situation is known as the “food versus fuel” debate. Further to this, first generation feedstocks are limited in their potential supply and their environmental credentials. Regrettably, a small proportion of the expansion of first generation biofuels comes as a result of deforestation to grow biofuel crops. This deforestation can also lead to soil erosion, increased levels of fertilizer application and emissions of nitrous oxides and chemical run-off. Second generation biofuels have the potential to overcome many of the negative issues associated with first generation biofuels. They are made from inedible feedstocks, which are often wastes from other processes. Typically these are lignocellulosic materials from wood and agricultural and other waste from agriculture (grown on arable land), urban centers or industry. They greatly increase the availability of the feedstock supply and have improved environmental credentials. However, second generation biofuels require large research and development costs to commercialize. Third generation biofuels have a distinct advantage over first and second generation biofuels in that the feedstock does not require arable land. The ability to use nonarable land, in principle, allows for potentially large enough volumes to replace the fossil fuel supply if other constraints could be overcome. Further, the use of wastes or byproducts from other industries is a common characteristic for third generation biofuels. It was originally envisaged that algae would use industrial wastewater to provide the nutrients for microorganisms with a high lipid content. Third generation biofuels have an inherent problem – if grown in open ponds, a large water supply is required with associated mass transfer challenges (e.g. high energy, high capital cost equipment). If grown in photo-bioreactors, the high water requirements can be overcome, but at present, capital costs are high.

1.4.1.1 Ligno-cellulosic wastes Ligno-cellulosic waste is produced by numerous industries: – Forestry sector waste. This includes material which comes from saw mill operations (e.g. bark and saw dust), logging (stumps) or plantation forestry maintenance such as thinnings (i.e. branches). This material is most commonly burned in a boiler for steam or electricity generation. – Pulp and paper industry black liquor. In order to make white papers, the industry takes woodchips, adds chemicals and dissolves the natural brown resin, lignin. In this process, the lignin is dissolved in the chemicals to form black liquor which must be burned as an economic and environmental measure. In doing so, the combustion of the lignin and incidental carbohydrate in the black liquor makes it a green energy source.

18 | 1 Matching the biomass to the bioproduct

–

–

Sugarcane industry waste. Sugarcane is harvested and crushed to extract the juice which contains the sugar. The fiber that is left behind is known as bagasse and the leaves left in the field are known as cane trash. Bagasse is most commonly used for steam and electricity generation in the factory but the electricity can also be exported for use in the community [6–10]. Other agricultural waste. The waste of many other agricultural industries is left in the field, such as straw. Some fruit and vegetable wastes are potential sources for bioenergy through anaerobic digestion.

The chemical composition of a ligno-cellulosic feedstock affects the potential bioproduct and the amount of energy produced. Compared to fossil fuels, biomass contains large amounts of moisture and oxygen which reduces the energy conversion efficiency. Apart from water, the most abundant chemical in ligno-cellulosic material is cellulose, which is a polymer of glucose, and has the formula (C6 H10 O5 )n . The proportion of cellulose in biomass can vary widely up to 90 % in cotton, although 20–60 % is more typical for wood and most agricultural crops. Cellulose can be broken down into its monomers by biological or chemical hydrolysis routes. However, accessibility of the cellulose depends on chemical composition and structure of the biomass. Hemicellulose is a random polymer of five and six carbon sugar monomers which is more prevalent in agricultural crops than in wood. The sugar monomers include glucose, mannose (C6 ), xylose, and arabinose (C5 ). Hemicellulose is readily broken down under acidic or basic conditions, as occurs in pulp production during paper manufacture. Lignin is a complex hydrophobic polymer composed of linked aromatic rings. Lignin provides the plant with a rigid structure and protects it from microbial attack. It inhibits fermentation and since it contains aromatic groups, when partially combusted it can produce toxic emissions. Its removal is necessary to allow fermentation to second generation biofuel and for the remaining cellulose and hemicellulose to be almost white and malleable, which is a requirement for white paper production. It is present in wood 25–40 %, and sugarcane 15–25 % [11] although other grasses contain almost no lignin (this is apparent by their color and lack of rigidity). There are three primary lignin monomers: p-Coumaryl, Coniferyl and Sinapyl alcohols (see Fig. 1.5). These monolignins differ in their reactivity due to the number of methoxy sites and each monolignin is present to varying extents in softwood, hardwood and grasses. OH

OH

CH3O

OCH3 OH

OH

OH

OCH3 OH

Fig. 1.5: Coumaryl, coniferyl, and sinapyl lignin [12].

1.4 Sample bioprocesses

|

19

1.4.1.2 Urban center waste The main potential feedstocks from urban centers are: – Municipal solid waste. Although this material contains significant amounts of plastics from fossil fuels, it has a high proportion of ligno-cellulosic content. This occurs due to the high amount of cardboard packaging society uses. Developed countries typically use over 100 kg of paper per capita each year. – Recycled fiber. The lignin content of this feedstock depends on whether it is generated from office waste (white papers) or household waste which includes boxes, which still contains significant amounts of lignin. – Sewage contains a very high organic content on a dry basis, although it has very high water content which is eliminated in many processing routes as conversion efficiency would be too low.

1.4.1.3 Regional considerations The availability of a feedstock depends on a variety of factors which must be assessed on a case by case basis. An approach to measuring the potential availability of feedstock is provided by Kosinkova et al. [13]. Kosinkova et al. used Australia, which was investigated as being a microcosm to demonstrate some of these issues as shown in Fig. 1.6. These considerations include [14, 15]: – Local population size. Is the region a significant urban center or an agricultural area? – The climate. Is the temperature better suited to forestry, agricultural fiber or a tropical crop such as sugarcane?

Sugarcane region

Large urban centres

Forestry region

Fig. 1.6: Distribution of bioenergy feedstocks in Australia.

20 | 1 Matching the biomass to the bioproduct

– – –

Availability of water. Agriculture and algal cultures benefit from an abundance of water supply. Logistical and political considerations. How does the waste management supply chain operate? Availability of local infrastructure to export material. E.g. electricity transmission lines, ports, roads, railways.

1.4.1.4 Processes and energy bioproducts The feedstocks are matched to the bioproducts by the conversion processes.

Combustion to produce electricity The oldest biomass to energy conversion process is simple combustion. Fiber is a form of hydrocarbon, and so burns in the presence of oxygen provided there is an ignition source. This is the basis for traditional household cooking and heating. Industrially, biomass is fed into the furnace of a large industrial boiler to generate steam and electricity via a turbine. The generalized formula for combustion is: Fuel + oxygen + nitrogen → water + carbon dioxide + nitrogen (+ carbon monoxide) , where carbon monoxide is produced as a result of incomplete combustion.

Anaerobic digestion to produce biogas Microorganisms break biomass down in the absence of oxygen in a series of steps which produce methane and carbon dioxide. Lignin is a problem for the initial degradation step, and so it is not suitable for wood. Apart from this, the process can be used for grasses, sewage, delignified waste (e.g. waste paper), abattoir waste, some household waste, fats and oils. The process can be tailored to the moisture content of the feedstock and multiple feedstocks can sometimes be used.

Fermentation to produce fuel ethanol Ethanol can be used as a liquid fuel and is particularly well suited for blending with petroleum gasoline. The conversion of sugars (sucrose, glucose) to ethanol involves converting glucose into ethanol and carbon dioxide through the action of microorganisms, namely C6 H12 O6 → 2C2 H5 OH + 2CO2 .

1.4 Sample bioprocesses

| 21

Transesterification to produce biodiesel Vegetable and animal oils consist of triglycerides which are reacted with methanol or ethanol in the presence of an acid or base catalyst in order to produce biodiesels. Glycerol is a byproduct of this reaction. The conversion to fatty acid methyl esters (i.e. FAME) is necessary to reduce the viscosity and improve atomization and combustion efficiency. O CH2 — O — CR1

O CH — O — CR2

R1 — C — O — R

+

3ROH

Catalyst

O CH2 — O — CR3

CH2 — OH

R2 — C — O — R

+

CH — OH

CH2 — OH

R3 — C — O — R

Thermochemical conversion to produce electricity or liquid fuel This is a family of routes which involve subjecting biomass to elevated temperatures in a tightly controlled oxygen environment (Fig. 1.7). Gasification occurs with a limited amount of oxygen present. Carbon monoxide and possibly carbon dioxide and carbon are produced, but these molecules react with water to produce hydrogen. The high temperatures result in high conversion efficiencies. Apart from hydrogen, the syngas can be used to produce methanol or liquid fuels (diesel) by the Fischer–Tropsch process [16].

C+H2O → H2+CO CO+H2O ↔ CO2+H2

Liquefaction

Pressure, bar

1000

100

“Syngas”

10

Gasification Pyrolysis

1 0

200

400 600 800 Temperature, °C

Fig. 1.7: Thermochemical conversion pathways.

1000

1200

2󸀠 second generation technology.

×

×

1󸀠 first generation technology;

×

×

Waste cooking oil and abattoir waste Microalgae

×

Transesterification (biodiesel)

×

×, 2󸀠 2󸀠

×, 1󸀠

× 1󸀠

Fermentation (fuel ethanol)

Vegetable oils

×

Sewage

Oils and fats

×

Municipal solid waste

Urban waste

× × ×

Anaerobic digestion (biogas)

Sugarcane bagasse Forestry waste Black liquor

Starch-based feedstocks (e.g. maize)

Sugar-rich feedstocks (e.g. sugarcane molasses, juice)

Combustion (electricity)

Lignin containing

Sugars and starch

Feedstock

Tab. 1.5: Matching biomass to bioenergy processes and bioproducts.

×, thermochemical liquefaction

×

×

×, gasification

× ×

Cheaper conversion pathways available

Cheaper conversion pathways available

Thermochemical conversion (to power, biochar, and/or oils)

22 | 1 Matching the biomass to the bioproduct

1.4 Sample bioprocesses

|

23

Pyrolysis occurs at more modest temperatures in the absence of oxygen and water. The conditions can be varied to target different products but generally bio-oil, solid char and volatile gases (e.g. methane) are produced. Flash pyrolysis, whereby the biomass is heated within just a few seconds, achieves better yields than slow pyrolysis. Liquefaction occurs at lower temperatures but high pressures are required. Liquefaction results in increased oil yields compared to pyrolysis, but the main advantage is that the process is highly tolerant of water and no pre-drying is needed, unlike in pyrolysis. The main disadvantage is that the equipment tolerant of these high temperatures and pressures is expensive. Table 1.5 shows typical matching of the feedstock to the process and bioenergy product. Sugars and starches are well suited for producing fuel ethanol, while combustion is best suited for drier lignified biomass. However, lignified biomass can be used to produce ethanol via second generation technology or converted via thermochemical processing. Urban wastes and oils are well suited to anaerobic digestion and thermochemical processing but oils are also used for biodiesel.

1.4.2 Nutraceutical example: Chondroitin Chondroitin sulfate is an important structural component of cartilage and provides resistance to compression. Chondroitin sulfate is used as a supplement for treatment of osteoarthritis and is believed to help draw water and nutrients into the cartilage, keeping it spongy and healthy. Chondroitin is a naturally occurring biomolecule found in the body and is a sulfated glycosaminoglycan (GAG) attached to proteins as part of a proteoglycan (hence the need to remove proteins during downstream processing via proteolysis). Made traditionally from shark cartilage, shark is considered a less sustainable source of chondroitin due to declining shark populations, hence bovine (cow) sources are more widely used. As a result, demand exists for “clean” sources of bovine chondroitin from a known and traceable source. An example process for the manufacture of chondroitin sulfate is presented in Fig. 1.8. It must be noted that the effectiveness of chondroitin and the combined use of glucosamine/chondroitin for osteoarthritis remains unclear; some studies have found chondroitin reduces pain more than a placebo, however several newer studies have found no improvement in pain with chondroitin; some evidence exists that chondroitin supplements slow cartilage breakdown or repair damaged cartilage from knee osteoarthritis [17]. Studies have found that glucosamine and chondroitin supplements may interact with the anticoagulant (blood-thinning) drug warfarin (Coumadin). Overall, studies have not shown any other serious side effects [18]. Typical doses of chondroitin sulfate for osteoarthritis are 200–400 mg two to three times daily or 1000–1200 mg as a single daily dose; when applied to the skin in a

24 | 1 Matching the biomass to the bioproduct

Biomass

Non-cartilage

Mechanical cleaning Cartilage

By-products processed into meal

Milling of cleaned cartilage

Enzyme proteolysis (e.g. alcalase, papain) Sediment Centrifugation Supernatant Ethanol recycle

Alkaline-ethanol proteolysis Sediment Re-dissolution and neutralization

Supernatant (recycle potential)

Ultrafiltration at 10–30 kDa cut-off Retentate Drying/Milling

Chondroitin sulfate powder, 99.5% purity

Fig. 1.8: Block diagram of a bioprocess for the manufacture of 99.5 % purity chondroitin sulfate.

cream for osteoarthritis typical doses are 50 mg chondroitin sulfate/g for up to 8 weeks [19]. Due to feedstock limitations, chondroitin sulfate sourced from shark cartilage is around five times the cost of chondroitin sulfate from bovine trachea. As an example of an alternative chondroitin containing product in the area of functional foods, dried whole beef tracheas containing high levels of naturally occurring glucosamine and chondroitin retail for approximately AU $ 51/kg retail finished product. In Fig. 1.9, the upstream process may be considered the “Producer” and “Processor” whilst the downstream process is represented by the “Value adder” (concentra-

1.4 Sample bioprocesses

| 25

tion into an impure bulk), and “Manufacturer” (purification). The retailer may then package the final purified product. As can be seen, the profit after tax (PAT) for the upstream sections is negligible whilst for the value adder PAT equates to 36 %, for the manufacturer 9 % and for the retailer 16 %. Chondroitin sulphate food grade value chain 1100.00

$AU per kg of retail finished product

1000.00 900.00

$1063 Purchase SG&A Depreciation PAT

COGS Marketing Tax

$175

800.00

Tax

$47

Marketing

$65 $24

PAT

Depre c

700.00 600.00

$352

SG&A

500.00 400.00 $75

$324

300.00 $213

200.00

$10 $5 $4

$76

$2

$12

Producer

Processor

$21 $32

Additional COGS

$324

$16

100.00 0.00

$28 $9 $54

$40

$213

$16

Value Adder Manufacturer

Retailer

Fig. 1.9: Graph showing the costs for producing chondroitin sulfate at various stages in the value chain. SG&A: Selling, General and Administrative Expenses. COGS: Cost of Goods Sold: the costs of making the products to be sold later. PAT: Profit after tax [20].

1.4.3 Fermentation example: Fermentation products from ligno-cellulose Given the abundance of ligno-cellulose and its availability as a co-product from industrial processes such as sugar milling, it is an attractive potential carbon source for industrial fermentations to produce a range of fuels and industrial chemicals. The majority of existing industrial microbial fermentation processes use glucose derived from corn starch or sucrose from sugarcane (in the form of molasses or cane juice) as the carbon source. To use ligno-cellulose as the feedstock, it is necessary to break down the constituent polymers to fermentable sugars, such as the conversion of cellulose to glucose [21, 22]. The challenge in the approach to use ligno-cellulosic materials arises from the recalcitrance of the feedstock. A major role of ligno-cellulose in nature is to provide mechanical strength to plant materials and resist degradation, meaning that significant time and energy is usually required to break it down. Further, once degraded, the economics of the fermentation processes would be maximized if all

26 | 1 Matching the biomass to the bioproduct

product streams (C5 and C6 sugars as well as lignin) could be utilized to generate products. To use all available sources of carbon means that multiple or highly adaptable processes are required (for example using microbes that can utilize both C5 and C6 sugars simultaneously) [23]. The overall process for the production of fermentation products from lignocellulose usually first involves a pretreatment step to mechanically and thermochemically degrade the material and separate the fibers, as well as degrade some carbohydrate polymers. This step converts the fibrous dry material to a liquid form where the remaining constituent polymers are accessible to enzymes for the second step of saccharification to generate fermentable sugars such as glucose or xylose. These sugars are then available as a carbon source for microbial fermentation to generate the desired end product chemical that must then be isolated and purified. Various purification and fractionation steps may be required during the entire process to isolate various constituents depending on the specific carbon source required for the fermentation and the need to remove microbial inhibitors such as furfural and phenolic compounds that may be produced during pretreatment [24]. The improvement of the overall process is an ongoing area of research and numerous options are being examined to increase process efficiency and reduce costs. The aims and goals of this research include reducing the temperature and overall severity of the pretreatment process to reduce the required energy, cost and levels of production of inhibitory molecules; the combination of enzymatic saccharification and fermentation into one step; the utilization of microorganisms that can both produce carbohydrase enzymes and ferment the resulting sugars to products in a so called consolidated bioprocess; and the use of microorganisms that can utilize both C5 and C6 sugars to produce the desired fermentation product.

1.4.3.1 Novel pretreatment strategies Typical pretreatment strategies usually employ a mechanical milling step to reduce particle size followed by a chemical modification step using dilute acid or alkaline hydrolysis along with further mechanical processing such as steam explosion [24]. Processes such as dilute acid hydrolysis typically employ temperatures from 120 to 210 °C and lead to significant hydrolysis of xylan to xylose as well as making cellulose available for enzymatic saccharification. Often such pretreatments lead to the formation of unwanted compounds that can inhibit the subsequent fermentation [24]. One approach to reduce the severity of the pretreatment and therefore reduce both energy costs and byproduct formation is to use co-solvents. Ionic liquids have been investigated for this purpose [25, 26] as well as acidified glycerol [27, 28], where increased enzymatic digestibility has been observed with increasing concentrations of glycerol. An ammonia-based process to expand the fiber in a processed termed AFEX™ has also been developed (for example see [29–31]). A direct comparison of dilute acid, ionic liquid and AFEX™ pretreatment methods was performed using corn stover [32].

1.4 Sample bioprocesses

|

27

The study found that similar ethanol production metabolic yields were observed for all three methods although the required enzyme combinations varied considerably to achieve the yields and the need for additional nutrient supplementation, which was not required after AFEX™ pretreatment. AFEX™ is being developed and commercialized by the not-for-profit company MBI in collaboration with Michigan State University. The technology has been tested in collaboration with Deinove using specific Deinococcus bacteria in a simultaneous saccharification and fermentation process that achieved high levels of utilization of the available sugars [33].

1.4.3.2 Use of specific ligno-cellulose fractions Once glucose can be generated from cellulose, it should be compatible with existing fermentation processes that utilize glucose derived from conventional sources, depending on the presence of inhibitory compounds. Industrial fermentations based on C5 sugars such as xylose and arabinose are not currently common in industry given that glucose or sucrose are generally more available feedstocks. Unlike cellulose that only generates glucose upon degradation, hemicellulose is a heterogeneous polymer that produces a range of both C5 and C6 (pentose and hexose) sugars and as such represents additional challenges for full utilization of the available sugars. Overall, to fully utilize the available sugars in ligno-cellulose, strains that can co-metabolize both C5 and C6 sugars are required. Ethanol production using Saccharomyces cerevisiae is one of the world’s oldest and largest bioprocesses with cellulosic ethanol production now entering commercial scale manufacturing [34]. Furthermore, this yeast is the basis for extensive metabolic engineering toward new chemical products [35], with some in commercial production (for example see the development of the farnesene process by the company Amyris). As such, S. cerevisiae has been a popular organism for the development of strains that can use C5 sugars such as xylose as well as glucose [36, 37]. These engineered strains must match the robustness of existing industrial ethanol strains and further, must ideally be tolerant of inhibitors and other constituents found in ligno-cellulosic hydrolysates, such as high levels of salt from pretreatment chemicals. For example, it has been shown that the presence of chloride and sulfate salts with sodium, potassium and ammonium cations reduced biomass growth, ethanol production and glucose consumption as well as reduced xylose utilization in the presence of sodium chloride in a S. cerevisiae strain capable of co-utilization of glucose and xylose [38]. An acetic acid reduction pathway to generate ethanol has also been integrated with C5 /C6 co-utilization in an S. cerevisiae strain [39]. Overall, S. cerevisiae has been extensively engineered to use a variety of carbon sources and generate a wide range of products [35] and remains a key platform organism in industrial biotechnology. Nevertheless, a range of other organisms have also been explored for similar aims, including Escherichia coli [40], Rhodococcus opacus [41] and Zymomonas mobilis [42].

28 | 1 Matching the biomass to the bioproduct

Lignin is a significant constituent of ligno-cellulose and presents numerous challenges for use associated with its natural role in providing protection and mechanical strength to biomass. It is also a heterogeneous and noncarbohydrate based polymer, meaning the breakdown products aren’t as readily utilized by microorganisms as glucose from cellulose. Lignin is generated at large scale in the pulp and paper industry and would be a major co-product from a future ligno-cellulosic ethanol industry. Lignin can be utilized by microorganisms for growth and metabolism offering a possible route to convert this challenging but highly available and cheap material to both biomass and chemical products. It has been known for some time that organisms such as white rot fungi are able to degrade ligno-cellulose using a suite of enzymes that break down lignin and enhance the availability of the carbohydrate polymer fractions to enzymatic saccharification [43]. The main classes of ligninolytic enzymes include lignin peroxidases, manganese-dependent peroxidases and laccases and these enzymes have been widely studied for the biocatalytic degradation of lignin [44, 45]. These enzymes can be isolated and used as biocatalysts to degrade and modify lignin although a collection of different enzymes are usually required in concert to achieve degradation. The use of enzymes has so far mostly been targeted at enhancing the pretreatment of biomass to isolate cellulose for breakdown to glucose [46]. As well as using isolated enzymes to transform the lignin polymer and associated monomers, the use of whole cell microorganisms and the manipulation of in vivo metabolic pathways for the degradation and assimilation of lignin derived chemicals is emerging as an option to generate specific products [44]. Lignin associated monomers such as ferulic acid and vanillic acid can be metabolized in vivo to protocatachuic acid and on to beta-ketoadipic acid and acetyl-CoA for incorporation into the TCA cycle and other pathways such as for fatty acid and isoprenoid synthesis. This channeling or funneling of a variety of lignin derived chemicals towards acetyl-CoA has been proposed as a solution to the problem of heterogeneity of the lignin substrate [47]. In this example, a specific strain of the bacterium Pseudomonas putida that is naturally able to catabolize aromatic molecules was used. Both lignin model compounds and heterogeneous lignin derived material could be converted to medium chain length polyhydroxyalkanoates. The same microorganism has also been metabolically engineered to produce cis,cis-muconate, an adipic acid precursor, from both the lignin model compound p-coumaric acid and a biomass-derived lignin stream [48]. The yield obtained on the model compound in fed-batch fermentations was 13.5 g/l and over 15 times what was achieved in shake flask cultures. Using the lignin stream in shake flask cultures produced 0.7 g/l of product at a molar yield of 67 % from the p-coumarate and ferulate substrates that were detected in the substrate. This work represents an encouraging first step towards further development of the process in bioreactors and additional improvements as both the lignin stream and microbial degradation metabolic pathways are further understood and optimized. An assessment of fourteen bacteria, including the P. putida strain described above, that secrete ligninolytic enzymes was performed to assess the formation of

1.4 Sample bioprocesses

|

29

molecules that could be used for industrial applications such as fuels, chemicals and materials [49]. This work provided information on particular species that could not only break down lignin but also metabolize the breakdown products to biomass and compounds such as polyhydroxyalkanoates, particularly in nitrogen limited conditions.

1.4.3.3 Consolidated bioprocessing As seen above for the consolidated bioprocessing of lignin directly to products using a single organism and process, the same concept can be applied to the processing of the cellulose and hemicellulose biomass fractions. There are two general strategies for the generation of microbial strains that can both break down ligno-cellulose to sugars and then convert them to chemical products. Such organisms generally do not exist in nature and so one must either endow the ability to produce cellulolytic enzymes on an existing production strain [50] or engineer a strain that already produces cellulolytic enzymes to be able to generate the product of interest [51]. There are many examples in the academic literature of where consolidated bioprocessing has been demonstrated, but commercial reality remains to be achieved due to cost, yield, efficiency and scalability issues in addition to the challenge of heterologous expression of cellulase enzymes [52]. An important and immediate application is the conversion of cellulose to glucose for the subsequent production of ethanol [53– 55]. The production of ethanol in this way builds on the existing and extensive ethanol biofuel industry. Making an established product such as ethanol is less risky than developing both cellulose degradation and engineering new molecule production at the same time. The relatively low selling price of ethanol as a fuel compared to producing higher value commodity or specialty chemicals means that commercial success for this concept remains challenging however. For reasons related to the co-utilization of hexose and pentose sugars, the utilization of hemicellulose in a consolidated bioprocess represents an additional challenge over and above the conversion of cellulose [56]. As well as requiring an organism that can produce relevant hydrolase enzymes (such as xylanases) and generate an industrial chemical product, the organism must also readily metabolize C5 and C6 sugars. The susceptibility of hemicellulose to degradation during acidic pretreatment steps, however, may alleviate the need for extensive enzymatic degradation. Nevertheless, the conversion of hemicellulose has been reported. One example involves the generation of succinic acid in an engineered strain of E. coli [57]. In this case, endoxylanases and xylosidases were selected for their suitability in a consolidated bioprocess, for example to have good activity at the ideal growth temperature of the bacterium. Enzyme production levels were optimized but secretion was found to be a limiting factor. Integration of the xylan utilization capability into an E. coli succinate production strain enabled the production of this industrially important chemical from xylan.

30 | 1 Matching the biomass to the bioproduct

1.4.3.4 Commercial applications With an established and large-scale ethanol biofuel industry, particularly in the USA and Brazil using corn and sugarcane feedstocks respectively, there has been significant industrial focus on generating commercial value from the ligno-cellulosic co-streams such as corn stover and bagasse. The large volumes of ligno-cellulosic biomass that could be available from these industries and others (including forestry) coupled with the lack of competition with human or animal food make this feedstock attractive despite the challenges of accessing the fermentable sugars. To this end the world’s major enzyme producing companies and research scientists backed by substantial government support have made significant progress in the development of improved cellulose enzymes. Along with pretreatment and process innovations, the first large-scale cellulosic ethanol production facilities have emerged. These facilities include Beta Renewables plant at Crescentino in Italy that has the potential to process 270 000 tons per year of biomass and generate up to 60 000 tons of ethanol. The first cellulosic ethanol facility in the USA was established in Iowa in 2014 by the POET-DSM joint venture [58]. It has the capacity to process 770 tons of biomass per day (281 050 tons per year operating 365 days per year) and generate 20–25 million gallons (60 000–75 000 tons) of ethanol per year. There exists significant scope for expansion of both the number and size of cellulosic ethanol production facilities once the technology and costs are fully established and proven. In 2015 there were 195 fuel ethanol plants with a mean production capability of 75 million gallons per year per plant according to U.S. Energy Information Administration statistics [59]. In Brazil GranBio is using sugarcane straw and bagasse at a facility in Alagoas and has the capacity to generate 82 million liters (65 000 tons) of ethanol per year. GranBio has also established a partnership with the chemical company Rhodia around using similar approaches for the production of n-butanol [60]. Research and development towards the production of 1,4-butanediol from ligno-cellulose has also been disclosed by the company Genomatica [61]. This work involved engineering E. coli for co-fermentation of C5 and C6 sugars, strain engineering towards 1,4-butanediol production from this feedstock and development of a fermentation process. A range of different hydrolysates from different processes and biomass types were also tested. Using a concentrated (700 g/l monomeric sugar) and clean hydrolysate, a 1,4-butanediol titer of 119 g/l and productivity approaching 2.5 g/l/h could be achieved. Overall, ligno-cellulosic biomass represents a significant resource for the future manufacture of liquid fuels and biochemical. The recalcitrant and heterogeneous nature of the material along with variations in composition between and within plant species raises many challenges for its economic and scalable use. These challenges also raise many opportunities for advancements in plant biotechnology for new biomass types, improved pretreatment technologies and the development of new microbial strains and fermentation processes to utilize as much of the available biomass and generate chemical products in the most efficient and cost effective manner.

1.4 Sample bioprocesses

|

31

1.4.4 Biopharmaceutical example: Monoclonal antibodies The rise of biopharmaceuticals has been unprecedented since the turn of the 21st century. Today biopharmaceuticals constitute around 20 % of the global pharmaceutical market, having a growth rate above 8 % per annum which is double that of chemical pharmaceuticals. Current biopharmaceutical revenues are estimated to be $ 163 billion [62]. Overall, biopharmaceuticals are the highest value biomolecules being produced from industrial processes but they also have a higher cost of manufacture than any other class of industrial biomolecules. Manufacturing costs are comparatively high for several reasons: facility outlay costs, regulatory compliance, lengthy processing times, low yields, expensive raw materials, costs of skilled staff and short market life cycles (e.g. competition from generics). In the case of the facility outlay, the costs to build a biopharmaceutical manufacturing facility start at $ 200–500 million and can take up to five years to complete depending on the nature of the facility and the biomolecules to be manufactured within the facility, whilst comparable chemical pharmaceutical facilities cost in the order of $ 30–100 million. The potential for disruptive process innovation in the biopharmaceutical sector is very high. However, the limitations are the cost-of-entry to develop and deliver a disruptive process which fits into existing facility designs, meets project budgets, project timelines and complies with regulatory requirements for validation, characterization and manufacture. Because of these limitations there is a constant cycle of reassessment of processes and investment in exploring new technology within the existing manufacturing landscape. Overall, process advancements have proven to be largely conservative and predominantly driven by project requirements to meet production needs and product quality. This has especially been the case for mammalian cell derived biopharmaceuticals which represent the highest cost biomolecules currently manufactured within the biopharmaceutical sector. For mammalian cell derived biopharmaceuticals the main innovations have come from:

Improvements to cell lines through metabolic engineering Metabolic engineering can improve consistency of fermentation, reduce media complexity and improve product yields and quality. For example, engineering the inclusion of glutamine synthetase allows a condensation reaction of glutamate and ammonia to form glutamine. For some cell lines – such as murine myeloma cells, there is no intrinsic glutamine synthetase and so engineering the cell line with glutamine biosynthetic capacity removes the challenges of glutamine supplementation to the media such as glutamine depletion/limitation or deamidation post-formulation of the media or during the course of fermentation. Such cell line developments produce clones with higher specific productivities for antibodies (exceeding 50 pg/cell/day) compared to the undeveloped base cell lines (less than 10 pg/cell/day).

32 | 1 Matching the biomass to the bioproduct

Molecular biology improvements to the DNA vector Improvements to the vector allow greater consistency of insertion into the host genome, thereby producing more stable clones and better transcription initiation. Stability is particularly important for mammalian cell lines such as murine and hamster cell lines as a large percentage of the genome (around 30 %) is composed of transposable elements that potentially can rearrange and insert into different regions of the genome. Industrial cell lines are qualified and validated to be stable for the duration of the manufacturing run (from vial thaw, through seed train expansion, to beyond the cell age seen at the end of the production cycle). Depending on the nature of the process, this can be up to 120 days where a cell line must remain consistent in terms of stability and productivity.

Process analytical technologies (PATs) PATs allow novel real-time monitoring of fermentation metrics and metabolism that provide greater elements of control and understanding of upstream processes. Greater uptake is now occurring for PATs that are proving to be fit-for-purpose for biopharmaceutical manufacture with the most pre-eminent being near infrared spectroscopy. Regulators are now setting in place guidelines for the use of PATs with a strong desire to see the uptake and implementation of PATs to improve consistency and quality of manufacture.

Biomass media and feed-media development Media and feedstocks are typically highly complex formulations and always provide the greatest increase in yield and quality metrics in simple batch and fed-batch fermentations. Due to regulatory constraints against the use of animal-derived materials (which may contain biological hazards such as bacteria, viruses, prions and endotoxins), industry has a preference for protein-free media with optimized concentrations of amino acids, salts, co-factors, vitamins and lipids, despite the high costs associated with Good Manufacturing Practice (GMP)-certified formulations. Typical yield increases from first generating a single stable clone to selection of the media and optimizing a feed strategy for batch fermentation are between two to tenfold for biopharmaceuticals. For fed-batch antibody processes of 10–21 days duration, titers in the range of 1–5 mg/l are common, but yields as high as 10–15 g/l have been reported. The advantages of optimized media include consistency of upstream biomass generation, the biopharmaceutical is safer (less contamination from difficult-to-purify animal products), the quality is more easily planned and controlled in the final product (referred to as quality by design or QBD) and the productivity of the biomass is higher. The focus of media development is to maximize the total viable cell density for as long as possible, which leads to significant biomass increases in fed-batch cultures. In conjunction with high specific productivity from engineered clones, economically

1.4 Sample bioprocesses |

33

viable processes can be established from relatively small bioreactor volumes. Optimized media will improve specific and volumetric yields as well as “time yields” (e.g. g product/hour of fermentation), a performance metric of interest for facilities that have capacity or fermentation volume limitations.

Single-use bioreactors (SUBs) The advent of SUBs represents a novel, disruptive technology for the generation of upstream biomass to produce biopharmaceuticals. The single-use bioreactor technology utilizes a sterilized disposable plastic bag as the bioreactor vessel which is then held in a temperature controlled jacketed support frame. SUB systems are complete, including all mixing, aeration and exhaust requirements (i.e. impellers, spargers and exhaust filters are integral to the sterilized disposable bags). The SUB systems are expensive – nowadays approximating the costs of cheaply sourced steel, but are considered to be economical when factoring all aspects of the pharmaceutical upstream process. This includes: – Production time considerations – sterilization using steam-in-place systems are not required and so turnaround times are improved – Batch loss and facility down-time due to contamination is greatly reduced – Flexibly designed facilities, which can be open-plan as SUB systems can be portable within a facility and therefore accommodate upstream and downstream changes or whole process differences for different biopharmaceuticals manufactured within the same facility. This decreases the costs of building a rigid facility which can be expensive to modify for evolving process requirements, and – Because the scales of the SUB technology are relatively small (usually 50 to 2000 l); the technology is more congenial for expansion by scaling-out rather than scaling-up, which simplifies operational expansion as the systems are the same and the complexities of different equipment at different scales are eliminated The SUB systems have the potential to be disruptive technology for the expensive and intensive biopharmaceutical workflow by allowing small operations and biotech start-ups to operate in niche areas and compete with large pharma at a greatly reduced cost. An example of niche manufacturing is monoclonal antibodies developed to Phase I levels by start-up pharmaceutical companies using wave-induced motion bioreactors. Wave-induced motion bioreactors were first described by Singh [63] and utilize the disposable, flexible and sterile plastic bag (cellbag) with controlled headspace gassing supported on a rocking thermo-regulated platform. The rocking platform is responsible for a distinctive undulating wave movement created in the bioreactor cellbag. This different, low-shear, mass and energy transfer mechanism ensures good off-bottom

34 | 1 Matching the biomass to the bioproduct

suspension, efficient nutrient distribution and a high oxygen transfer from a large surface mixing area. Cell growth is controlled by adjusting the rocking rate, the rocking angle, the fill level, headspace aeration rate and headspace gas composition. Customized and more complex systems have been described that control cell growth with nutrient feed controls, integrated perfusion and online monitoring-feedback controls of dissolved oxygen (DO) and pH (controlled via O2, CO2 and base addition) [64, 65]. Wave-induced motion bioreactors can be used as a process bioreactor on a range of different cells for both stable and transient systems, or as a unit operation for biomass expansion for seeding process bioreactors and high-density cell banking. Though simple in design and operation, the performance characteristics have been demonstrated to be distinct – but similar – to more complex stirred tank bioreactors [66]. Simple changes such as the percentage of CO2 in the headspace can control the pH of the media or potentially alter the pH outside of a normal operating range. The hydrodynamic flow (mixing and shear behavior) is not necessarily the same in different sized cellbags or with different media volumes. However, there is generally a high mass transfer coefficient (kL a), that is unusually steady over a broad range of operational parameters (the kLa is reported to be as high as 20–30/h at operation ranges from 20–30 rpm, 7.5° [67]). The kL a can be simply manipulated by changing the rocking angle, rocking rate and, to a very superficial extent due to the large surface area of exchange, the headspace gassing. As such, most operational conditions for the wave-induced motion bioreactors can be achieved in a short amount of time and manipulated simply; for example, it is known that subtle changes in the percentage of CO2 in the headspace can elicit the effect of either stripping or enriching the pCO2 as desired to influence the media pH in a reasonable timeframe of just over 10 min for a 50 l bioreactor without any other control system for the pH [68]. Furthermore, scaling can be readily achieved when considering the media:cellbag ratio and adjusting for differences in the hydrodynamic flow changes for different sized cellbags and different media volumes. The “kL a fixing” to match different scales typically requires manipulation of the integral power density via aeration and/or agitation rates, with smaller media volumes having higher kL a values and requiring less power for kL a fixing. A smaller media to cellbag ratio is predicted to have higher kL a values and more readily become influenced by the power terms (rocking angles and rocking rates) and gas velocity (headspace gassing). Successful and simple scaling therefore allows a smaller fermentation model to be established to reduce the costs associated with development and characterization. For monoclonal antibody production, a small-scale model system using a waveinduced motion bioreactor of 10–50 l media can be used as an affordable development platform to establish a process to then scale to kL a fixing anywhere up to 500 l. Using a perfusion system, a continuous cell culture process can be established at a very modest scale whereby biomass is retained in the wave-induced motion bioreactor, while new culture media is continuously added at approximately one (1) bioreactor volume

1.4 Sample bioprocesses

| 35

per day and culture supernatant is removed at the same rate to keep the bioreactor volume constant. In this respect a 10 l scale-down perfusion model can be used to economically develop a Phase I process. When scaled to a modest 100 l wave-induced motion bioreactor, the operation in perfusion mode from day 7 to day 21 (14 day perfusion) is equivalent to operating in batch or fed-batch at a scale of 1400 l. If a conservative 1 g/l volumetric yield is achieved from such a process and assuming a low 30 % yield from downstream unit operations, such a process could produce approximately 400 g of monoclonal antibodies and therefore be suitable to initiate Phase I investigations at a greatly reduced cost.

1.4.5 Novel material example: Cellulose nanofibers 1.4.5.1 Manufacture and properties of cellulose nanofibers Cellulose is the most abundant natural polymer on earth. The fundamental building block of cellulose in all ligno-cellulosic materials is of crystallites of 4–5 nm in diameter. Thus, given the degree of polymerization of cellulose, the natural organization of cellulose in all ligno-cellulosic material is as cellulose nanofibers [69]. Cellulose nanomaterials are produced by breaking down ligno-cellulosic material, either partially or fully, into the constituent nanofibers. The terminology around cellulose nanofibers is still in flux. Broadly the materials may be divided into two classes, based on crystallinity and aspect ratio. Cellulose nanocrystals or nanocrystalline cellulose are treated with acid to hydrolyze the cellulose. Mechanical treatment, to break down the structure produces 5–10 nm diameter, low aspect ratio cylinders of crystalline cellulose. Microfibrillated cellulose (MFC) or cellulose nanofibers are high aspect ratio material, often only partially separated, produced by combining mechanical treatment, typically with a chemical pretreatment to reduce the energy required for separation. Cellulose nanofibers are typically more heterogeneous than nanocrystalline cellulose, both in the average fiber diameter for a given batch and the distribution of fiber diameters [70]. In the rest of this brief discussion, the focus will be on the applications of cellulose nanofibers. A characteristic of this class of material is that it can be made from almost any ligno-cellulosic material including hardwoods and softwoods [71] and many different non-wood fibers [72] and grasses [73], including a variety of agricultural residues. It is generally considered that for optimal separation of fibers into cellulose nanofibers, all or most of the lignin should be removed prior to processing [74]. Many different methods can be used to separate the starting material into cellulose nanofibers, including homogenization [75], grinding [76], ball milling [77] and refining. The common elements are applying shear force to a low consistency (often around 1 %) suspension. Without appropriate pretreatment, the energy consumption required to fully separate the starting material into nanofibers in a homogenizer is very large, perhaps up to 20 000 kWh/t, thus leading researchers to investigate more

36 | 1 Matching the biomass to the bioproduct

energy efficient ways to treat the fibers or developing pretreatments to reduce energy consumption by enhancing separation efficiency. Currently available pretreatments include enzymatic treatment [75], carboxymethylation or TEMPO assisted oxidation [78]. Pretreatment typically also produces more uniform fibers.

1.4.5.2 Applications of cellulose nanofibers Applications of cellulose nanofibers can be broadly divided into additive applications and sheet and/or film applications. Cellulose nanofibers have been widely investigated as a reinforcing fiber in composites. Matrices investigated include petrochemical derived matrix materials such as polypropylene [79]. However, the use of bioderived materials such thermoplastic starch [80] or PLA [81] to make all biocomposites, seems to be the most promising approach. The fibers do not require additional treatment to compatibilize them with the biopolymer matrix and the final product is renewable and biodegradable. Adding only 5 wt.% of cellulose nanofibers to PLA, increased tensile strength and elastic modulus by almost 25 % [81]. Cellulose nanofibers can also be added directly to the matrix in twin-screw extrusion [81], where wood pulp fibers cannot, as they are three orders of magnitude larger. Cellulose nanofibers also show considerable promise as an additive in conventional papermaking. Cellulose nanofiber addition improves both wet and dry strength and increases the ratio of wet to dry strength [82, 83], while reducing permeability and increasing sheet resistance to drainage [84]. Sheet strength was doubled [83] by using a combination 50–100 mg/g of nanofibers and 5 mg/g of PAE, a polymeric strength additive. The improvement in mechanical strength is because the aspect ratio of the cellulose nanofibers is much higher [85] than all hardwood and many softwood fibers. The strength of short fiber nonwovens such as paper typically increases with aspect ratio and is only controlled by fiber length if the diameter is constant. Higher aspect ratio increases the efficiency of stress transfer within the network, by reducing the importance of load build up from the ends of the fibers [86]. Sheet or film applications include as membranes, as barrier layers and for substrates for flexible electronics [87], which will not otherwise be discussed here. Standalone sheets are formed from suspensions either by filtration [70, 88, 89] or casting [90]. Casting is extremely time consuming and is not likely to be a commercially viable solution for large-scale production. Filtration can be time consuming due to the dimensions of the nanofibers [89]. The small size of the fibers means that the filtration conditions must be carefully selected in order to retain the nanofibers on the filter media. In addition, as the filter mat forms, the small size of the pores in the structure reduces the flow through the filter mat. Sheet preparation time can be significantly reduced by using high solids content to minimize water to be removed, combined with filters with large mesh openings to minimize filter resistance [70, 88]. The addition

1.4 Sample bioprocesses |

37

of small amounts of polyelectrolytes such as CPAM and PAE will also further improve the drainage speed [91, 92]. Methods of application of nanofiber layers onto substrates include filtration [93], spray coating [94] and contact coating [95]. One of the most interesting commercial possibilities for the use of cellulose nanofibers is as a barrier layer. Paper as a packaging material has significant weaknesses. The fibers are hydrophilic and make a network with high porosity composed of large micron sized pores, which pose little barrier to water or gas transport. Up until now, paper based packaging requiring excellent barrier properties has been made as polymer composites or laminates, with the paper providing the stiffness and the polymer materials or layers controlling the permeability. However, such composite materials are difficult to recycle. Cellulose nanofiber barrier layers are effective, firstly, because the nanofibers typically form a denser network than sheets made from conventional wood fibers. Sheet densities ranging from 780 kg/m3 to 1400 kg/m3 have been reported, in comparison to the density of cellulose of approximately 1500 kg/m3 . Secondly, the flow rate through a structure of a given porosity, for a given pressure drop, will scale with the size of the pores, which in turn scale with the fiber diameter. Reducing the fiber diameter by three orders of magnitude and increasing the network density, reduces the network permeability to close to that of typical polymer laminate layers, while remaining fully compatible and recyclable with the packaging. Because the cellulose is hydrophilic, cellulose nanofiber layers typically have poorer resistance to water vapor transport. Cellulose nanofiber layers of 35 grams per square meter (gsm) had water vapor transmission rate (WVTR) of 215 g/m2/day/m compared to low density polyethylene (LDPE) which had WVTR of 100 g/m2/day/m [90]. The WVTR could be reduced to below that of the LDPE film by either adding mineral filler or an additional coating of cooked starch [90], neither of which would be expected to affect the recyclability of the film. In contrast, the oxygen permeability is more than two orders of magnitude lower than either low or high density polyethylene [96] and is only slightly worse than specialist barrier polymers such as EVOH. Treatment with TEMPO mediated oxidation to produce uniform, very low diameter nanofibers has been shown to produce excellent oxygen barrier properties even with a film thickness of only 0.4 µm [95]. The wide range of application modes and the ability to tailor the barrier performance suggests products are likely to be commercialized in the near term. Applications of cellulose nanofiber sheets as ultrafiltration membranes have been demonstrated [93]. The experiments prepared membranes by filtering nanofiber suspensions through a support layer, followed by pressing and conventional drying. Membrane layer grammage ranged from 5 to 30 gsm, using commercial MFC, with some of the larger fibers removed and the layer density estimated from other studies is expected to be 780 kg/m3 [92]. Porosity and pore size were also tailored by adding silica nanoparticles. The results showed that MFC alone produced a porous structure with the pores too large for conventional ultrafiltration, but that addition of 10 % by weight of silica nanoparticles reduced both the average size and dispersion of the pore

38 | 1 Matching the biomass to the bioproduct

sizes, producing a membrane that performed adequately for higher end ultrafiltration applications, although the performance still needs further improvement compared to commercial membranes. The major possible application of the material is as an ultrafiltration membrane, separating based on size. Another application has recently been demonstrated in oil-water separation. Oilwater mixtures are extremely heterogeneous, ranging from fully or partially separated phases to dispersed emulsions. However, even well-emulsified mixtures will typically have a particle size in the micron range. So the pore size required for size separation is much higher than for ultrafiltration applications, and separation is often only gravity driven. As well as size exclusion, separation is also engineered by chemical compatibility, i.e. by using the hydrophilicity of the cellulose or by modifying the cellulose fibers to make them hydrophobic [97], oleophobic or oleophilic. Pore size has been increased compared to conventional sheets by freeze drying partially separated MFC, where the energy consumption in fibrillation was reduced so as to only partially separate the fibers [97]. This process has been shown to produce sheet materials with density as low as 0.002 kg/m3 [97]. This field of research is still in development. It has been the intent of this brief survey to highlight the potential of cellulose nanofibers as a renewable, recyclable nanomaterial, which can greatly extend the property space achievable with conventional renewable fiber materials.

1.5 Conclusion Dramatic increases in the demand for energy and products over the past decades with an associated requirement for lower greenhouse gas (GHG) emissions has led to the need to revisit the feedstocks and processes that society uses to generate power, fuels and goods. The consumption of fossil fuels has resulted in a number of negative environmental outcomes, including acid rain, water contamination and global warming. The challenge is to utilize low cost feedstocks to create products and energy in ways that are technically, environmentally and economically viable. Hence, there exists an urgent need for the development of innovative, flexible, efficient and low emissions technologies that utilize low cost feedstocks. Future works can utilize computer based process modelling to rapidly consider and optimize a huge range of feedstock-unit operation-product combinations for meeting the future needs of society. Such work will highlight knowledge gaps that need to be filled with regards to physical data, reaction kinetics, rates of reaction, plant design and quality/composition requirements for creating the highest value products for the lowest cost. For example, algae offer the opportunity to biosequester carbon via the creation of biochar from a pyrolysis process, however, may firstly require the creation of several higher value biomolecules to be economically viable. Conversely, the co-processing of low cost feedstocks such as bagasse, green wastes, and/or coal can provide feedstock reliability and economies of scale.

References

|

39

Bioprocess engineers can address the major challenges facing the development and application of new industries by: – Addressing the limited data on the kinetics of complex reactions involved in processing biomass and co-processing of mixed feedstocks – Higher capital and operating costs that represent obstacles for adopting these technologies – Restrictive and continually more constraining emissions to air requirements to ensure that industry is sustainable – Rapidly changing operating environments which will require industrial facilities to be flexible with regards to feedstocks due to dramatic changes to commodities prices, changes to regional and global climates, and extreme weather events – Life cycle analysis/quadruple bottom line analysis (economic, environmental, societal, GHG)

References [1] [2] [3] [4] [5]

[6] [7] [8]

[9]

[10] [11]

[12] [13]

Deloitte Touche Tohmatsu, Opportunities for the fermentation-based chemical industry. 2014. Data Monitor, Pharmaceuticals: Global Industry Guide. Research and Markets Publishing, Dublin, Ireland, 2010. Industry Experts. Biopharmaceuticals- A Global Market Overview. 2013. Transparency Market Research, Neutraceutical ingredients market. Albany, 2014. Chum H, Faaij A, Moreira J, Berndes G, Dhamija P, Dong H, Gabrielle B, Gosseng A, Lucht W, Mapako M, Masera Cerutti O, McIntyre T, Minowa T, Pingoud K. Bioenergy, in Edenhofer O et al., editors. IPCC Special Report on Renewable Energy Sources and Climate Change Mitigation, Cambridge University Press: Cambridge, 2011. Rainey TJ, Doherty B, Martinez DM, Brown R, Kelson NA. The effect of flocculants on the filtration properties of bagasse pulp, Tappi Journal, 2010 (May), 7–14. Rainey TJ, Doherty WOS, Martinez DM, Brown RJ, Kelson NA. Pressure filtration of bagasse pulp, Transport in Porous Media, 2010, 86(3), 737–751. Rainey TJ, O’Hara I, Mann AP, Bakir CH. Reducing bagasse dust emissions from stockpile operations by depithing, in Proceedings of the Australian Society for Sugarcane Technologists, 2012. Rainey TJ, O’Hara I, Mann AP, Bakir CH, Plaza F. Effect of depithing on the safety and environmental aspects of bagasse stockpiling, Process Safety and Environmental Protection, 2013, 91, 378–385. Rainey TJ. A comparison between highly depithed and conventionally depithed bagasse pulp, Appita Journal, 2012, 65(2), 178–183. Rainey TJ. A study into the permeability and compressibility properties of Australian bagasse pulp, in Faculty of Built Environment and Engineering, Queensland University of Technology: Brisbane, 2009. Doherty B, Rainey T. Bagasse fractionation by the soda process, in Proc. Aust. Soc. Sugar Cane Technol. Mackay, 2006. Kosinkova J, Doshi A, Maire J, Ristovski Z, Brown R, Rainey TJ. Measuring the regional availability of biomass for biofuels and the potential for microalgae, Renewable & Sustainable Energy Reviews, 2015, 49, 1271–1285.

40 | 1 Matching the biomass to the bioproduct

[14] Covey G, Rainey TJ, Shore D. A new opportunity to pulp bagasse in Australia, in Proc. Aust. Soc. Sugar Cane Technol. Mackay, 2006. [15] Rainey TJ, Covey G, Shore D. An analysis of Australian sugarcane regions for bagasse paper manufacture, International Sugar Journal, 2006, 108(1295), 640–644. [16] Ramirez J, Brown RJ, Rainey TJ A Review of Hydrothermal Liquefaction Bio-Crude Properties and Prospects for Upgrading to Transportation Fuels, Energies, 2015, 8, 6765–6794. [17] Arthritis Australia. Glucosamine and Chondroitin. 2014 [Accessed on 26 February 2016] http:// www.arthritisvic.org.au/Complementary-Therapies/Complementary-Therapies/Glucosamineand-Chondroitin. [18] National Center for Complementary and Integrative Health. Glucosamine and Chondroitin for Osteoarthritis: What You Need To Know 2014 [Accessed on 26 February 2016] https://nccih.nih.gov/health/glucosaminechondroitin. [19] U.S. National Library of Medicine. Chondroitin sulfate. 2015 [Accessed on 26 February 2016] http://www.nlm.nih.gov/medlineplus/druginfo/natural/744.html. [20] Greenwoods W, editor. Bioactive market study of beef, steep and goat meat: value chain analysis. M.a.L.A. Ltd, 2011. [21] Chundawat SPS, Beckham GT, Himmel ME, Dale BE. Deconstruction of lignocellulosic biomass to fuels and chemicals, Annual Review of Chemical and Biomolecular Engineering, 2011, 2(2), 121–145. [22] Blanch HW, Simmons BA, Klein-Marcuschamer D. Biomass deconstruction to sugars, Biotechnology Journal, 2011, 6(9), 1086–1102. [23] Kim SR, Ha SJ, Wei N, Oh EJ, Jin YS. Simultaneous co-fermentation of mixed sugars: a promising strategy for producing cellulosic ethanol, Trends in Biotechnology, 2012, 30(5), 274–282. [24] van der Pol EC, Bakker RR, Baets P, Eggink G. By-products resulting from lignocellulose pretreatment and their inhibitory effect on fermentations for (bio)chemicals and fuels, Appl Microbiol Biotechnol, 2014, 98(23), 9579–93. [25] Vancov T, Alston AS, Brown T, McIntosh S. Use of ionic liquids in converting lignocellulosic material to biofuels, Renewable Energy, 2012, 45, 1–6. [26] Klein-Marcuschamer D, Simmons BA, Blanch HW. Techno-economic analysis of a lignocellulosic ethanol biorefinery with ionic liquid pre-treatment, Biofuels Bioproducts & Biorefining-Biofpr, 2011, 5(5), 562–569. [27] Zhang Z, Wong HH, Albertson PL, Doherty WO, O’Hara IM. Laboratory and pilot scale pretreatment of sugarcane bagasse by acidified aqueous glycerol solutions, Bioresour Technol, 2013, 138, 14–21. [28] Zhang Z, Wong HH, Albertson PL, Harrison MD, Doherty WO, O’Hara IM. Effects of glycerol on enzymatic hydrolysis and ethanol production using sugarcane bagasse pretreated by acidified glycerol solution, Bioresour Technol, 2015, 192, 367–73. [29] Bals BD, Gunawan C, Moore J, Teymouri F, Dale BE. Enzymatic Hydrolysis of Pelletized AFEX (TM)-Treated Corn Stover at High Solid Loadings, Biotechnology and Bioengineering, 2014, 111(2), 264–271. [30] Jin MJ, Gunawan C, Balan V, Lau MW, Dale BE. Simultaneous saccharification and cofermentation (SSCF) of AFEX (TM) pretreated corn stover for ethanol production using commercial enzymes and Saccharomyces cerevisiae 424A(LNH-ST), Bioresource Technology, 2012, 110, 587–594. [31] Chundawat SPS, Bals B, Campbell T, Sousa L, Gao D, Jin M, Eranki P, Garlock R, Teymouri F, Balan V, Dale BE. Primer on Ammonia Fiber Expansion Pretreatment, in Aqueous Pretreatment of Plant Biomass for Biological and Chemical Conversion to Fuels and Chemicals, 2013, pp. 169–200.

References

|

41

[32] Uppugundla N, Sousa LD, Chundawat SPS, Yu XR, Simmons B, Singh S, Gao X.D, Kumar R, Wyman CE, Dale BE, Balan V. A comparative study of ethanol production using dilute acid, ionic liquid and AFEX (TM) pretreated corn stover, Biotechnology for Biofuels, 2014, 7. [33] Deinove. DEINOVE teams up with MBI, pioneer of AFEX technology, to evaluate its process on industrial biomass. 2014 [Accessed on 26 February 2016] http://www.deinove.com/en/ news/all-press-releases/deinove-teams-mbi-pioneer-afex-technology-evaluate-its-processindustrial-biomass. [34] Dionisi D, Anderson JA, Aulenta F, McCue A, Paton G. The potential of microbial processes for lignocellulosic biomass conversion to ethanol: a review, J. Chem. Technol. Biotechnol, 2015, 90, 366–383. [35] Borodina I, Nielsen J. Advances in metabolic engineering of yeast Saccharomyces cerevisiae for production of chemicals, Biotechnology Journal, 2014, 9, 609–620. [36] Weber C, et al. Trends and challenges in the microbial production of lignocellulosic bioalcohol fuels, Applied Microbiology and Biotechnology 2010, 87(4), 1303–1315. [37] Matsushika A, et al. Ethanol production from xylose in engineered Saccharomyces cerevisiae strains: current state and perspectives, Applied Microbiology and Biotechnology, 2009, 84(1), 37–53. [38] Casey E, Mosier NS, Adamec J, Stockdale Z, Ho N, Sedlak M. Effect of salts on the Cofermentation of glucose and xylose by a genetically engineered strain of Saccharomyces cerevisiae, Biotechnology for Biofuels, 2013, 6(83). [39] Wei N, Oh EJ, Million G, Cate JHD, Jin YS. Simultaneous utilization of cellobiose, xylose, and acetic acid from lignocellulosic biomass for biofuel production by an engineered yeast platform, Synthetic Biology, 2015, 4(6), 717–713. [40] Kim SM. Simultaneous utilization of glucose and xylose via novel mechanisms in engineered Escherichia coli, Metabolic Engineering, 2015, 30, 151–148. [41] Fei Q. High-cell-density cultivation of an engineered Rhodococcus opacus strain for lipid production via co-fermentation of glucose and xylose, Process Biochemistry, 2015, 50(4), 500–506. [42] Dunn KL, Rao CV. Expression of a xylose-specific transporter improves ethanol production by metabolically engineered Zymomonas mobilis, Appl Microbiol Biotechnol, 2014, 98(15), 6897–905. [43] Leonowicz A, Matuszewska A, Luterek J, Ziegenhagen D, Wojtas-Wasilewska M, Cho NS, Hofrichter M, Rogalski J. Biodegradation of lignin by white rot fungi, Fungal Genet Biol, 1999, 27(2–3), 175–85. [44] Bugg TDH, Rahmanpour R. Enzymatic conversion of lignin into renewable chemicals, Current Opinion in Chemical Biology, 2015, 29, 10–17. [45] Wong DWS. Structure and Action Mechanism of Ligninolytic Enzymes, Applied Biochemistry and Biotechnology, 2009, 157(2), 174–209. [46] Rico A, Rencoret J, del Rio JC, Martinez AT, Gutierrez A. Pretreatment with laccase and a phenolic mediator degrades lignin and enhances saccharification of Eucalyptus feedstock, Biotechnology for Biofuels, 2014, 7. [47] Linger JG, Vardon DR, Guarnieri MT, Karp EM, Hunsinger GB, Franden MA, Johnson CW, Chupka G, Strathmann TJ, Pienkos PT, Beckham GT. Lignin valorization through integrated biological funneling and chemical catalysis, Proceedings of the National Academy of Sciences of the United States of America, 2014, 111(33), 12013–12018. [48] Vardon DR, Franden MA, Johnson CW, Karp EM, Guarnieri MT, Linger JG, Salm MJ, Strathmann TJ, Beckham GT. Adipic acid production from lignin, Energy & Environmental Science, 2015, 8(2), 617–628.

42 | 1 Matching the biomass to the bioproduct

[49] Salvachua D, Karp EM, Nimlos CT, Vardon DR, Beckham GT. Towards lignin consolidated bioprocessing: simultaneous lignin depolymerization and product generation by bacteria, Green Chemistry, 2015. [50] den Haan R, et al. Progress and challenges in the engineering of non-cellulolytic microorganisms for consolidated bioprocessing, Current Opinion in Biotechnology, 2015, 33, 32–38. [51] Hasunuma T, Okazaki F, Okai N, Hara K.Y, Ishii J, Kondo A. A review of enzymes and microbes for lignocellulosic biorefinery and the possibility of their application to consolidated bioprocessing technology, Bioresour Technol, 2013, 135, 513–22. [52] Lambertz C, et al. Challenges and advances in the heterologous expression of cellulolytic enzymes: a review, Biotechnology for Biofuels, 2014, 7(1), 135. [53] Gholamreza SJ, Taherzadeh MJ. Advances in consolidated bioprocessing systems for bioethanol and butanol production from biomass: a comprehensive review, Biofuel Research Journal, 2015, 2(1), 152–195. [54] Hasunuma T, Kondo A. Development of yeast cell factories for consolidated bioprocessing of lignocellulose to bioethanol through cell surface engineering, Biotechnology Advances, 2012, 30(6), 1207–1218. [55] Hasunuma T, Kondo A. Consolidated bioprocessing and simultaneous saccharification and fermentation of lignocellulose to ethanol with thermotolerant yeast strains, Process Biochemistry, 2012, 47(9), 1287–1294. [56] Girio FM. Hemicelluloses for fuel ethanol: A review, Bioresource Technology, 2010, 101(13), 4775–4880. [57] Zheng Z, Chen T, Zhao M, Wang Z, Zhao X. Engineering Escherichia coli for succinate production from hemicellulose via consolidated bioprocessing, Microb Cell Fact, 2012, 11, 37. [58] POET-DSM. First commercial-scale cellulosic ethanol plant in the U.S. opens for business. 2014 [Accessed on 26 February 2016] http://www.dsm.com/corporate/media/informationcenternews/2014/09/29-14-first-commercial-scale-cellulosic-ethanol-plant-in-the-united-statesopen-for-business.html. [59] U.S. Energy Information Administration. U.S. Fuel Ethanol Plant Production Capacity. 2015 [Accessed on 26 February 2016] http://www.eia.gov/petroleum/ethanolcapacity/. [60] GranBio. GranBio-Rhodia. 2015 [Accessed on 26 February 2016] http://www.granbio.com.br/ en/conteudos/biochemicals/. [61] Burk M, Barton N, Trawick J. Development of an Integrated Biofuel and Chemical Refinery. Final report for Genomatica. [Accessed on 26 February 2016] http://www.genomatica.com/ _uploads/pdfs/Genomatica,BiomassAdvances,TechnicalPaper,June2015.pdf. [62] Otto R, Santagostino A, Schrader U. The beauty and the beast: A perspective on biopharmaceuticals, in Otto R, Santagostino A, Schrader U, editors. From Science to Operations Questions, Choices and Strategies for Success in Biopharma, McKinsey & Company, 2014, p. 10. [63] Singh V. Disposable bioreactor for cell culture using Wave-induced agitation, Cytotechnology, 1999, 30, 149–158. [64] Clincke M, Mölleryd C, Zhang Y, Lindskog E, Walsh K, Chotteau V. Study of a recombinant CHO cell line producing a monoclonal antibody by ATF or TFF external filter perfusion in a WAVE Bioreactor™, in 22nd European Society for Animal Cell Technology (ESACT) Meeting on Cell Based Technologies. Vienna, Austria, 2011, p. P105. [65] Clincke M, Mölleryd C, Zhang Y, Lindskog E, Walsh K, Chotteau V. Very High Density of CHO Cells in Perfusion by ATF or TFF in WAVE Bioreactor™. Part I. Effect of the Cell Density on the Process, Biotechnol. Prog, 2013, 29(3), 754–767. [66] Rodrigues ME, Costa A.R, Henriques M, Azeredo J, Oliveira R. Wave characterization for mammalian cell culture: residence time distribution, New Biotechnology, 2012, 29(3), 402–408.

References

|

43

[67] Tao Y, Shih J, Sinacore M, Ryll T, Yusuf-Makagiansar H. Development and Implementation of a Perfusion-Based High Cell Density Cell Banking Process, Biotechnol. Prog, 2011, 27(3), 924–929. [68] Yuk IH, Baskar D, Duffy PH, Hsiung J, Leung S, Lin AA. Overcoming challenges in WAVE Bioreactors without feedback controls for pH and dissolved oxygen, Biotechnol. Prog, 2011 27(5), 1397–406. [69] Isogai A. Wood nanocelluloses: fundamentals and applications as new bio-based nanomaterials, Journal of Wood Science, 2013, 1–11. [70] Zhang L, Batchelor W, Varanasi S, Tsuzuki T, Wang X. Effect of cellulose nanofiber dimensions on sheet forming through filtration, Cellulose, 2012, 19(2), 561–574. [71] Syverud K, Chinga-Carrasco G, Toledo J, Toledo PG. A comparative study of Eucalyptus and Pinus radiata pulp fibers as raw materials for production of cellulose nanofibrils, Carbohydrate Polymers, 2011, 84(3), 1033–1038. [72] Alemdar A, Sain M. Isolation and characterization of nanofibers from agricultural residues Wheat straw and soy hulls, Bioresource Technology, 2008, 99(6), 1664–1671. [73] Amiralian N, Annamalai PK, Memmott P, Schmidt S, Taran E, Martin D. Easily deconstructed, high aspect ratio cellulose nanofibers from Triodia pungens; an abundant grass of Australia’s arid zone, RSC Advances, 2015. [74] Spence KL, Venditti RA, Habibi Y, Rojas OJ, Pawlak JJ. The effect of chemical composition on microfibrillar cellulose films from wood pulps: Mechanical processing and physical properties, Bioresource Technology, 2010, 101(15), 5961–5968. [75] Henriksson M, Henriksson G, Berglund LA, Lindström T. An environmentally friendly method for enzyme-assisted preparation of microfibrillated cellulose (MFC) nanofibers, European Polymer Journal, 2007, 43(8), 3434–3441. [76] Abe K, Iwamoto S, Yano H. Obtaining cellulose nanofibers with a uniform width of 15 nm from wood, Biomacromolecules, 2007, 8(10), 3276–3278. [77] Zhang L, Tsuzuki T, Wang X. Preparation of cellulose nanofiber from softwood pulp by ball milling, Cellulose, 2015. [78] Saito T, Nishiyama Y, Putaux J.-L, Vignon M, Isogai A. Homogeneous suspensions of individualized microfibrils from TEMPO-catalyzed oxidation of native cellulose, Biomacromolecules, 2006 7(6), 1687–1691. [79] Suzuki K, Okumura H, Kitagawa K, Sato S, Nakagaito A, Yano H. Development of continuous process enabling nanofibrillation of pulp and melt compounding, Cellulose, 2013, 20(1), 201–210. [80] Hietala M, Mathew AP, Oksman K. Bionanocomposites of thermoplastic starch and cellulose nanofibers manufactured using twin-screw extrusion, European Polymer Journal, 2012. [81] Jonoobi M, Harun J, Mathew AP, Oksman K. Mechanical properties of cellulose nanofiber (CNF) reinforced polylactic acid (PLA) prepared by twin screw extrusion, Composites Science and Technology, 2010 70(12), 1742–1747. [82] Su J, Mosse WKJ, Sharman S, Batchelor WJ, Garnier G. Effect of tethered and free microfibrillated cellulose (MFC) on the properties of paper composites, Cellulose, 2013, 20, 1925-1935. [83] Ahola S, Österberg M, Laine J. Cellulose nanofibrils – Adsorption with poly(amideamine) epichlorohydrin studied by QCM-D and application as a paper strength additive, Cellulose, 2008, 15(2), 303–314. [84] Taipale T, Österberg M, Nykänen A, Ruokolainen J, Laine J. Effect of microfibrillated cellulose and fines on the drainage of kraft pulp suspension and paper strength, Cellulose, 2010, 17(5), 1005–1020. [85] Varanasi S, He R, Batchelor WJ. Estimation of cellulose nanofiber aspect ratio from measurements of fiber suspension gel point, Cellulose, 2013, 20(4), 1885–1896.

44 | 1 Matching the biomass to the bioproduct

[86] Batchelor WJ. An analytical solution for the load distribution along a fiber in a nonwoven network, Mechanics of Materials, 2008, 40(12), 975–981. [87] Zhu H, Fang Z, Preston C, Li Y, Hu L. Transparent paper: fabrications, properties, and device applications, Energy & Environmental Science, 2014, 7(1), 269–287. [88] Varanasi S, Batchelor WJ. Rapid preparation of cellulose nanofiber sheet, Cellulose, 2013, 20(1), 211–215. [89] Sehaqui H, Liu A, Zhou Q, Berglund LA. Fast preparation procedure for large, flat cellulose and cellulose/inorganic nanopaper structures, Biomacromolecules, 2010, 11(9), 2195–2198. [90] Spence KL, Venditti RA, Rojas OJ, Pawlak JJ, Hubbe MA. Water vapor barrier properties of coated and filled microfibrillated cellulose composite films, BioResources, 2011, 6(4), 4370–4388. [91] Raj PR, Varanasi S, Batchelor WJ, Garnier G. Effect of cationic polyacrylamide on the processing and properties of nanocellulose films, Journal of Colloid & Interface Science, 2015, 447, 113–119. [92] Varanasi S, Batchelor WJ, Superior non-woven sheet forming characteristics of low-density cationic polymer-cellulose nanofiber colloids, Cellulose, 2014, 21(5), 3541–3550. [93] Varanasi S, Low Z-X, Batchelor WJ. Cellulose nanofiber composite membranes – biodegradable and recyclable UF membranes, Chemical Engineering Journal, 2015 265, 138–146. [94] Beneventi D, Chaussy D, Curtil D, Zolin L, Gerbaldi C, Penazzi N. Highly porous paper loading with microfibrillated cellulose by spray coating on wet substrates, Industrial & Engineering Chemistry Research, 2014, 53(27), 10982–10989. [95] Fukuzumi H, Saito T, Wata T, Kumamoto Y, Isogai A. Transparent and high gas barrier films of cellulose nanofibers prepared by TEMPO-mediated oxidation, Biomacromolecules, 2009, 10(1), 162–165. [96] Syverud K, Stenius P. Strength and barrier properties of MFC films, Cellulose, 2009, 16, 75–85. [97] Wang S, Peng X, Zhong L, Tan J, Jing S, Cao X, Chen W, Liu C, Sun R. An ultralight, elastic, costeffective, and highly recyclable superabsorbent from microfibrillated cellulose fibers for oil spillage cleanup, Journal of Materials Chemistry A, 2015, 3(16), 8772–8781.

Meng Liang, Xiaowei Zhou, and Chunping Xu

2 Systems biology in biofuel 2.1 Introduction Biotechnology has played an important role in society, life, and economics since ancient times. Application of microorganisms in biotechnology demonstrates the use of cell factories for the production of a wide variety of chemicals used for fuels, commodities, specialty chemicals, polymers, and drugs [1, 2]. Current global problems, including environmental pollution, global warming, and energy security, have led to an increasing interest in renewable production of fuels and other chemicals currently derived from petroleum, especially the production of biofuels from lignocellulosic biomass [3]. The International Energy Agency (IEA) has predicated a more than 10-fold increase in biofuel demand from 2010 to 2050, resulting in a total demand of more than 30 exajoules [4]. The currently availability of biofuels, such as bioethanol and plantoil derived biodiesel, is not sufficient to meet current demand. Additionally, due to its physical properties, such as low energy density and high hygroscopy, bioethanol is not an ideal substitute for gasoline. Advanced biofuels with superior fuel and operational properties, including a diversity of C4 -C5 chain alcohols, biodiesel (fatty acid ethyl esters (FAEEs) and farnesane) and jet fuels (alkanes, olefins, and terpenes), are not naturally produced in preferred cell factories, e.g. Escherichia coli and Saccharomyces cerevisiae. Therefore, novel pathways have to be introduced to enable microorganisms to produce new and advanced biofuels. Since the advent of recombinant DNA technology, the production of desired natural products has been greatly enhanced. Especially in the past three to four decades, metabolic engineering has been developed as a powerful approach to increase production of these useful chemicals through directed genetic engineering [2, 5]. In the early stages of metabolic engineering, the development of this technology was mainly focused on improving the product yield and range of natural and unnatural chemicals, as well as the substrate utilization rates by targeted modification of metabolic pathways in host microorganisms. With recent advances, metabolic engineering has been applied to promote the design and development of novel cell factories [6–10], which, through its requirement of a cell-wide understanding and modification of microorganism metabolism, contributed to the growth of synthetic biology. To promote the production of biofuels using microorganisms, a significant modification of intracellular metabolism is required [1, 11, 12]. The ability for cell-wide metabolic engineering and synthetic biology, however, requires a system-level understanding of cellular metabolism. With the development of omics techniques, metabolic toolkits are needed to modify or build large numbers of metabolic pathways to produce chemicals in a feasible manner [13, 14]. The application of such techniques has severe consequences on metabolisms of the host

46 | 2 Systems biology in biofuel

strains. Therefore, a deep understanding of cellular metabolism is necessary for the design of strategies. With the advancement of omics techniques, thousands of parameters can now be monitored simultaneously [15, 16], making diagnosing and fixing metabolic engineering problems feasible. To analyze the vast amounts of data, systemwide models of various aspects on genetic regulation and metabolism as well as corresponding computational tools are needed, which introduce systems biology [17, 18]. Governments and companies show high interest in the production of biofuels. This indicates a successful and fast development of biofuel production in the near future. This review will cover the recent developments in biofuels from a technological point of view supported by systems biology. The development of biodiesel, jet fuels and butanol will be discussed in more detail, especially with regard to how tools from systems biology may advance the implementation of metabolic engineering processes and promote the development of novel biorefinery processes.

2.2 The importance of systems biology In traditional molecular biology approaches, only a few metabolic engineering modifications would be evaluated simultaneously, making a system-level perspective difficult to obtain. Understanding the metabolisms in a system-wide view, however, is strictly necessary for the success of metabolic modification strategies. With the application of systems biology, cellular phenotypes can be analyzed in more detailed and in a whole view, further leading to the improvement of cell-factory design through detailed metabolic modeling. Today, by introducing different omics techniques for cell factories, a large number of cellular components can be analyzed [18, 19]. The omics technologies show their deep influence on the development of metabolic engineering strategies. For example, the complex regulatory pathways involved in controlling metabolism, such as the protein kinase in yeast [20, 21], identification of transcriptional influence to metabolic fluxes in yeast and E. coli [22, 23] and the interaction among protein kinases in yeast [9]. With the integration of the omics techniques and system-level modeling approaches, metabolic engineering strategies can be drawn out rationally (Fig. 2.1). In the following parts, the omics techniques and system-level modeling are brief discussed.

2.2.1 Genomics The study of genomics provides information for acquiring and interpreting genomic sequence data. A staggering amount of genomics now exists for hundreds of different microbial and nonmicrobial systems, providing information on mutation, adaption, and growth conditions based on cell-wide data [24]. With the recent development in techniques and reduction in sequencing costs it is possible to use genome sequencing

2.2 The importance of systems biology |

47

Systems biology Omics techniques

Metabolic modeling Xylose

Glucose

GLYC3P

Metabolomics Proteomics Transciptomics Genomics

G6P

H2O

F6P

Xu5P

FBP

GAP

S7P

E4P

F6P

DHAP

6PGC

GAP 13BPG

GLYC

Interaction between omics and modeling

6PGL

NH4

3PG

PPi

2PG

HCO3

Pi CO2

EtOH

AC CIT

AMP ADP

ALLPHN

AcALD

AcCoA

O2

ICIT

OAA

ATP

MAL

2 CytCox

CoQH2

2 CytCred

CoQ

FUM

Xylitol

Reactions

R5P XYLU

Cellmass

PEP PYR

Ru5P

L-GLN

Urea

Metabolites

Fluxomics

CO2

0 1 0 0 0 ⁞

0 0 1 –1 0 ⁞

0 –1 0 1 0 ⁞

0 0 0 0 0 ⁞

1 0 0 0 –1 ⁞

… … … … … …

S-matrix

GLO

AKG L-GLU SuccCoA Succ 4AB SucSA

Metabolic engineering strategies Fig. 2.1: The derivation of metabolic engineering strategies from systems biology. The interaction between omics data and metabolic modeling would promote the understanding of the target organisms and lead to the implementation of rational metabolic engineering strategies.

and RNA sequencing to obtain information at the genome level as well as improved annotation and expression profiling [25]. The technology has also enabled rapid profiling of microbial diversity [26] and gene function [27] from complex, environmental samples. The genome-sequencing technique has been used to identify driving mutations among adaptively evolved strains in studies of E. coli [28] and of yeast [29], which proved its power. The demand to obtain and process the large amount of data obtained from transcriptional array studies have raised the need for entirely new disciplines combining bioinformatics and computational biology [17], and therefore required the development of protocols for rigorous reporting [30]. However, with the possibility of identifying genomic mutations, genome data alone has difficulty to identify governing mutations without help from additional omics technologies for detailed phenotypic analysis [29]. This kind of difficulty is usually caused by the appearance of silent mutations resulted from adaptive evolution. Therefore, it is desired to integrate the genomic data with other omics data [31].

2.2.2 Transcriptomics Genome-wide transcription analysis is the most widely used omics technology in metabolic engineering, which has been applied to various industrially relevant microorganisms, such as yeast [32], E. coli [33, 34] and Aspergilli [35]. The value of transcription analysis lies in its scale, which is genome-wide, and therefore can be integrated directly with genome-scale metabolic models via different approaches [36– 38]. Transcription and protein expression data correlated relatively well for detailed pathways [17], however, the correlation is usually poor for cell-wide analyses [39, 40],

48 | 2 Systems biology in biofuel

which indicates regulatory processes have a higher importance than transcriptional control. A study on mRNA synthesis and degradation in S. cerevisiae showed that both of them can influence protein levels significantly [41]. Meanwhile, stressors could also affect mRNA stability [42, 43]. With the integration of proteomic data, in addition to transcription, post-translational modification [44], protein localization [45, 46], and protein-protein interactions showed importance to cellular responses to stress [47]. Functional analysis of S. cerevisiae under particular stress conditions showed that candidate genes essential for growth did not show significant change [25], which emphasizes the importance of responses beyond differential expression.

2.2.3 Proteomics Just as mentioned above in transcriptomics, proteomics plays an importance role in metabolism and has advanced significantly in the past decades [48]. As the primary approach to capture regulation of cellular response beyond mRNA level, proteomics provides important information for functional genomics studies. Traditionally, proteomics are measured with two-dimensional electrophoresis [49] while nowadays high-throughput techniques via mass spectrometry have become the foundation for research. With the development of strategies for protein identification [49, 50] and workflow for protein abundance quantification [51], proteomics provides wideranging information such as composition, function and location. Among the various techniques, the iTRAQ technique, which can simultaneously label samples with up to eight conditions via isobaric tags [52], has been applied to detect protein changes [53] and to determine members of protein complexes [54]. Besides the progress of measurement approaches, computational analysis has to be improved to reduce artifacts in information and false positives [55–57]. With decades’ development, the standardized approaches that can report and handle proteomics data statistically soundly are still evolving [58, 59].

2.2.4 Metabolomics In order to wholly understand the enzyme activity and substrate turnover, monitoring transcriptome and proteome alone do not provide enough information. Metabolomics are providing true measurements of the cellular response to stress and manipulation [9, 60], and there has been a huge increase both in fields of application and the number of metabolites that can be measured [61, 62]. Methods have been developed for metabolome analysis of microorganisms [63–66], but key challenges still exists in sampling and extraction [67, 68]. Meanwhile, the absence of reliable and standardized databases, which should contain all of the existing spectral information and allow correct identification of unknown detected metabolites, is still one of the

2.2 The importance of systems biology

|

49

problems affecting metabolomics research. Additionally, the data on metabolite levels from different laboratories based on various experimental methods are not quantitatively consistent while the relative levels are comparable [69].

2.2.5 Fluxomics Additional to the above omics, fluxomics has shown its usages in studies of a range of various industrial microoganisms [70, 71]. The fluxome integrates information on cellular processes, and shows the unique phenotypic characteristics of the metabolisms of the cells [72]. The metabolome can be captured by flux analysis through its functional interactions with the environment and the genome [73]. Therefore, the fluxome is always accompanied by knowledge of the metabolome. With current techniques, it is still not that easy to measure intracellular fluxes. Thus, computational methods are combined together with experimental methods, in which the most reliable approaches are based on isotope-labeled precursors of metabolic pathways, mainly 13 C-labeled substrates [32, 74]. Via metabolomics analytical platforms, the concentration and isotopomeric distribution (or labeling pattern) of the labeled metabolites can be determined [72, 75]. However, the estimation of fluxes based on tracer metabolomics data requires a priori knowledge of possible distributions of the tracer used within the network, i.e. the structure and components of the network. Therefore, the lack of information on reactions and metabolites might lead to erroneous results, which means that a further understanding of other omics information is very important to the accuracy of fluxomes [76].

2.2.6 Computational Methods Metabolic modeling has been an integral part of metabolic engineering since its formation. With recent developments, by integration of these system-wide data from omics technology, mathematical modeling has been applied to design cell factories, which is another key contribution of systems biology to metabolic engineering [77]. Integrated models have been applied in many aspects for metabolic engineering, such as identifying essential genes [78], transcriptional elements [16], regulatory circuits [79], and stress response [15]. The models, therefore, provide valuable information about intracellular activities such as reaction fluxes and may lead to the discovery of the as yet unrevealed reasons preventing optimal production [80]. To integrate the omics data with metabolic models effectively, various computation methods have been developed to try to address issues with data integration [81]. Valuable tools have been published with open sources, which has made it much easier for researchers to analyze, compare, and mine genomics data [82–84]. The development of the tools has recently been reviewed extensively [14, 36, 85–87], while model repositories and

50 | 2 Systems biology in biofuel

on-line modeling tools [88, 89] will significantly promote the application of these models in the field of industrial biotechnology.

2.3 Applicability of systems biology in biofuels In order to combine native and heterologous genes in metabolic engineering [90], understanding how the incorporation of an engineered exogenous pathway perturbs the host system is important for overcoming pathway bottlenecks. Reaching desired production levels of metabolites requires a significant amount of pathway optimization. The application of systems biology in strain development for biofuel production is critical to identify potential bottlenecks and reveal detrimental side effects [21]. With the development of computational optimization approaches, in silico simulation may prove a product of interest possible to be produced in vivo with rational metabolic engineering strategies. Therefore, the undesired side products can be depressed while the carbon flux towards the target product can be maximized. This approach has been applied in the metabolic engineering improvement of bio-ethanol production from E. coli [5]. In the following parts, applications of metabolic engineering strategies based on integration of omics techniques and systems-level modeling are briefly reviewed in the production of advanced biofuels, specifically, biodiesels, jet fuels and biobutanol.

2.3.1 Biodiesels The development of biodiesel techniques, just as other biofuels, focuses on a consistent, scalable and renewable commodity supply and a cost-efficient production based on the requirements of a sustainable society. To reach the target, metabolic engineering strategies, derived from understanding metabolism with the help of systems biology, have to be applied to construct a single cell factory. For biodiesel synthesis, usually, two metabolic pathways, the lipid and isoprenoid pathways, are employed [91]. Several research groups have investigated the metabolic engineering approaches to improving FAEE production in E. coli as well as the fatty acid metabolism [92–96]. By introducing cytosolic expression of thioesterase (󸀠 TesA), modulation of β-oxidation (by overexpression of faaD and deletion of fadE) and introduction of wax ester synthase/diacylglycerol acyltransferase (WS/DGAT) from Acinetobacter sp. ADP1, an engineered E. coli strain has been constructed which produced 400 mg/l FAEEs on glucose and 2 % ethanol in 2 days [96]. To further promote the production of FAEEs, the alcohol synthesis pathway (pdc and adhB) from Zymomonas mobilis and a second copy of atfA and acetyl-CoA ligase from S. cerevisiae were further introduced, which increased the production of biodiesel up to 674 mg/l based on glucose [96]. By employing different thioesterases, the distribution of product chain lengths was under control. Additional overexpression of acetyl-CoA carboxylase (accBACD) led to a final concentration of

2.3 Applicability of systems biology in biofuels

| 51

922 mg/l FAEEs in a scaled up fed-batch fermentation [97]. The main products from the fermentation consisted primarily of ethyl palmitate, ethyl oleate, ethyl myristate and ethyl palmitoleate. In order to further improve the production, the pyruvate dehydrogenase complex (aceEF) could be overexpressed. Meanwhile, the interruption of pathways to lactate, formate and acetate by deleting corresponding genes ldhA, pflB, poxB and ackA respectively, as well as phospholipid pathway through deleting gpsA and plsB, have been applied to reduce byproduct formation. Besides the work on E. coli, S. cerevisiae has also been modified to study its capacities on the production of biodiesels [98]. The removal of genes DGA1, LRO1, ARE1, and ARE2 led to the loss of capacity of storage lipids (TAGs and steryl esters) synthesis. Low FAEEs could be produced by the strain with the introduction of WS/DGAT [99]. An additional approach has also been applied by recycling glycerol into biodiesels by yeast. During the process, the introduced glycerol was converted into ethanol and was further converted into FAEEs by external addition of fatty acids and introduction of endogenous transesterification reaction catalyzed by WS/DGAT from Acinebacter sp. ADP1. The modification of the metabolic pathways led to a minimized glycerol secretion with 0.52 g/l FAEEs production rate and 17 g/l glycerol consumption rate [100]. Farnesane production has also been studied systematically, which requires introducing farnesene synthase to convert farnesyl pyrophosphate (FPP) to farnesene and further from farnesene to farnesane. By overexpressing the mevalonate pathway and fusing heterologous FPP synthase (ispA) and the apple α-farnesene synthase gene (wFS), E. coli produced a final concentration of 380 mg/l α-farnesene [101]. Besides, by engineering the mevalonate pathway and introducing bisabolene synthase from Abies grandis Ag1, both E. coli and S. cerevisiae reached a production of bisabolene higher than 900 mg/l, which is a precursor of bisabolane. The latter has been identified as an alternative fuel compound [102].

2.3.2 Jet fuels For a jet fuel, the must-have characteristics include low freezing point (–40 °C), high energy density (> 53.4 MJ/l) and comparable net heat combustion [103]. The commonly recognized candidates include n-alkanes, n-alkenes (n-olefins) and terpene compounds (pinene, sabinene and terpinene) [104]. Among the above chemicals, alkanes/alkenes have received further attention. By applying systems biology approaches, several pathways in bacteria for producing long-chain (C10 < x < C20 ) and very-long-chain (> C20 ) alkanes have been discovered. Comparison of cyanobacteria with and without alkanes product capacity led to the discovery of two genes that encoded enzymes for producing fatty aldehydes and converting further to alkanes by removing the carbonyl carbon [105]. The two encoded enzymes have been proven to be acyl-ACP reductase and aldehyde deformylase respectively [106, 107]. The functions of the two enzymes have been proven by introducing into E. coli, which led

52 | 2 Systems biology in biofuel

to a production of up to 300 mg/l of pentadecane, pentadecene, and heptadecene. Further adaptation of metabolisms by the introduction of the enzymes include increasing NADPH consumption and balanced protein reductive system for aldehyde deformylase. Meanwhile, acyl-CoA reductase can also produce fatty aldehydes (and theoretically alkanes) [108]. FFA can be converted to acyl-CoA synthetases and further into fatty alcohols, which have better price and performance than diesel [109]. A second route for alkane production, which was identified in a Jeotgalicoccus sp. by a reverse genetic approach, is catalyzed by P450 enzyme OleTJE [110]. When heterologously expressed in E. coli, the protein led to production of 1-pentadecene and 1,10-heptadecadiene as well as 1-heptadecene with addition of stearic acid (18 : 0). The results suggest that OleTJE can utilize both acyl-ACPs and FFA [109]. In addition to the two pathways mentioned above, Shewanella oneidensis can head-to-head condensate fatty acids to long-chain alkenes (C23 –C33 ) [111, 112]. Besides alkanes, olefins have been produced in several bacteria, which can be blended with diesel as well as be used as chemical building blocks. Synechococcus sp. PCC7002 produces 1-nonadecene and 1,14-nonadecadience [113]. Ols was found to be responsible for the synthesis of the olefins. Sulfation is the activation center for the protein. Its current titers are still very low while engineering the gene might lead to a set of pathways for producing a wide range of olefins [109].

2.3.3 Biobutanol As the natural butanol producer, Clostridium acetobutylicum has been recognized as an industrial cell factory for 1-butanol production and has been extensively studied [114]. Recently, progress on genetic engineering of Clostridium has been made [115]. Metabolic engineering strategies have been applied to eliminate byproducts (such as ethanol, acetone) [116, 117], to improve the oxygen tolerance and to reduce growth inhibition [118, 119]. While the study of Clostridium revealed its metabolic mechanisms, the heterologous strains were also built based on the results. Two different approaches have been applied to E. coli. The first approach is to modify E. coli’s highly active amino acid biosynthetic pathway, which led to a 1-butanol production of 0.5 g/l [90]. Another method is to introduce the 1-butanol pathway from C. acetobutylicum [120]. The host strain produced a similar amount of 1-butanol as the first method. In order to further promote 1-butanol production, several pathways have been introduced into E. coli. With the introduction of enzyme Crt and AdhE2 from the 1-butanol pathway of C. acetobutylicum, PhaA and Hbd from Ralstonia eutropha and Ter from Treponema denticola as well as the overexpression of aceEF to increase the acetyl-CoA pool, the constructed E. coli showed an evaluated metabolic system improvement and produced 1500-fold increase of 1-butanol titer from 3 mg/l to 5 g/l [121, 122]. Additional modification of metabolic pathways by blocking NADH reducing pathways (i.e. deleting frdBC,

2.4 Conclusions and outlook

| 53

ldhA, adhE, and pta to block succinate, lactate, ethanol and acetate pathways respectively), implementing an NADH drain in 1-butanol pathway through the enzymes from C. acetobutylicum (inserting Hbd, Crt and AdhE) and introducing highly active endogenous acetyle-CoA acyltransferase (AtoB) and irreversible trans-enoyl-CoA reductase (Ter) from T. denticola, led to a 15 g/l 1-butanol production. In situ removal of product from fermentation broth further doubled the yield to 30 g/l [123], which indicated that the product toxicity-level has to be kept low for proper cell functioning [124, 125]. Meanwhile, S. cerevisiae is also considered as a better host strain for 1-butanol than E. coli due to its higher butanol tolerance and long application in industrial ethanol production [126, 127]. However, with systems biology approaches, in silico simulations suggested lower butanol and propanol yields in yeast compared to E. coli, which was partially due to the limited flexibility of the central metabolism [128]. By introducing various enzymes to construct corresponding butanol production pathways from E. coli, Clostridium beijerinckii and Ralstonia eutropha, the constructed S. cerevisiae produced a 1-butnaol concentration of 2.5 mg/l [129]. Further modifications, such as increasing the acetyl-CoA pool by overexpression of pyruvate dehydrogenase multienzyme complex (lpdA, ace, aceF) and downregulating pyruvate decarboxylase (PDC), did not show much better results [111]. Therefore, more modification based on systems biology is still needed to construct engineering strains with high 1-butanol production.

2.4 Conclusions and outlook Cell factories for the production of biofuels, including biodiesel [93] and isobutanol production [90], have made substantial progress in recent years. Further improvements on the metabolism of the microorganisms, however, are still needed to meet the industrial requirements for advanced biofuels in terms of yield, titer and productivity. A systems-level understanding and analysis of metabolism will reveal strategies that can help to improve the productivities as well as other important factors such as tolerance towards the product of interest and the ability to use complex feedstocks derived from biomass. Many strategies for metabolic engineering have been proposed based on recent cell-wide studies to improve various aspects of biofuel production with the developments on omics techniques [130–132]. Systems biology is necessary to fully understand the regulation mechanisms of the underlying metabolic network and the response to the overproduction of molecules for the host in order to deduce pathway bottlenecks and optimize the design of cell factories. With the advancement of systems biology methods and tools, the application of systems biology approaches in metabolic engineering will become a key technology for the future creation of robust and efficient cell factories for industrial applications.

54 | 2 Systems biology in biofuel

References [1] [2] [3] [4] [5] [6] [7]

[8] [9] [10] [11] [12]

[13] [14] [15] [16] [17] [18] [19] [20]

[21] [22]

Fortman JL, Chhabra S, Mukhopadhyay A, et al. Biofuel alternatives to ethanol: pumping the microbial well, Trends Biotechnol, 2008, 26(7), 375–81. Chang MCY, Keasling JD. Production of isoprenoid pharmaceuticals by engineered microbes, Nat Chem Biol, 2006, 2(12), 674–81. Herrera S. Bonkers about biofuels, Nat Biotechnol, 2006, 24(7), 755–60. Fairley P. Introduction: Next generation biofuels, Nature, 2011, 474(7352), S2–5. Dien BS, Cotta MA, Jeffries TW. Bacteria engineered for fuel ethanol production: current status, Appl Microbiol Biotechnol, 2003, 63(3), 258–66. Alper H, Stephanopoulos G. Global transcription machinery engineering: a new approach for improving cellular phenotype, Metab Eng, 2007, 9(3), 258–67. Ostergaard S, Olsson L, Johnston M, Nielsen J. Increasing galactose consumption by Saccharomyces cerevisiae through metabolic engineering of the GAL gene regulatory network, Nat Biotechnol, 2000, 18(12), 1283–6. Farmer WR, Liao JC. Improving lycopene production in Escherichia coli by engineering metabolic control, Nat Biotechnol, 2000, 18(5), 533–7. Zhang J, Vaga S, Chumnanpuen P, et al. Mapping the interaction of Snf1 with TORC1 in Saccharomyces cerevisiae, Mol Syst Biol, 2011, 7, 545. Gibson DG. Gene and genome construction in yeast, Curr Protoc Mol Biol, 2011, Chapter 3, Unit3.22. Wackett LP. Biomass to fuels via microbial transformations, Curr Opin Chem Biol, 2008, 12(2), 187–93. Blanch HW, Adams PD, Andrews-Cramer KM, Frommer WB, Simmons BA, Keasling JD. Addressing the need for alternative transportation fuels: the Joint BioEnergy Institute, ACS Chem Biol, 2008, 3(1), 17–20. Nielsen J, Jewett MC. Impact of systems biology on metabolic engineering of Saccharomyces cerevisiae, FEMS Yeast Res, 2008, 8(1), 122–31. Tyo KEJ, Kocharin K, Nielsen J. Toward design-based engineering of industrial microbes, Curr Opin Microbiol, 2010, 13(3), 255–62. Ishii N, Nakahigashi K, Baba T, et al. Multiple high-throughput analyses monitor the response of E. coli to perturbations, Science, 2007, 316(5824), 593–7. Bonneau R, Facciotti MT, Reiss DJ, et al. A predictive model for transcriptional control of physiology in a free living cell, Cell, 2007, 131(7), 1354–65. Ideker T, Galitski T, Hood L. A new approach to decoding life: systems biology, Annu Rev Genomics Hum Genet, 2001, 2, 343–72. Sánchez BJ, Nielsen J. Genome scale models of yeast: towards standardized evaluation and consistent omic integration, Integr Biol (Camb), 2015, 7(8), 846–58. Fritzsch FSO, Dusny C, Frick O, Schmid A. Single-cell analysis in biotechnology, systems biology, and biocatalysis, Annu Rev Chem Biomol Eng, 2012, 3(1), 129–55. Usaite R, Jewett MC, Oliveira AP, Yates JR, Olsson L, Nielsen J. Reconstruction of the yeast Snf1 kinase regulatory network reveals its role as a global energy regulator, Mol Syst Biol, 2009, 5, 319. Jones JA, Toparlak ÖD, Koffas MAG. Metabolic pathway balancing and its role in the production of biofuels and chemicals, Curr Opin Biotechnol, 2015, 33, 52–9. Fendt S-M, Oliveira AP, Christen S, Picotti P, Dechant RC, Sauer U. Unraveling conditiondependent networks of transcription factors that control metabolic pathway activity in yeast, Mol Syst Biol, 2010, 6(432), 432.

References

[23]

[24] [25] [26]

[27] [28]

[29]

[30] [31] [32]

[33] [34] [35]

[36]

[37] [38]

[39]

[40] [41]

| 55

Haverkorn van Rijsewijk BRB, Nanchen A, Nallet S, Kleijn RJ, Sauer U. Large-scale 13C-flux analysis reveals distinct transcriptional control of respiratory and fermentative metabolism in Escherichia coli, Mol Syst Biol, 2011, 7(1), 477. Warner JR, Patnaik R, Gill RT. Genomics enabled approaches in strain engineering, Curr Opin Microbiol, 2009, 12(3), 223–30. Giaever G, Chu AM, Ni L, et al. Functional profiling of the Saccharomyces cerevisiae genome, Nature, 2002, 418(6896), 387–91. DeSantis TZ, Brodie EL, Moberg JP, Zubieta IX, Piceno YM, Andersen GL. High-density universal 16S rRNA microarray analysis reveals broader diversity than typical clone library when sampling the environment, Microb Ecol, 2007, 53(3), 371–83. He Z, Gentry TJ, Schadt CW, et al. GeoChip: a comprehensive microarray for investigating biogeochemical, ecological and environmental processes, ISME J, 2007, 1(1), 67–77. Conrad TM, Joyce AR, Applebee MK, et al. Whole-genome resequencing of Escherichia coli K-12 MG1655 undergoing short-term laboratory evolution in lactate minimal media reveals flexible selection of adaptive mutations, Genome Biol, 2009, 10(10), R118. Hong K-K, Vongsangnak W, Vemuri GN, Nielsen J. Unravelling evolutionary strategies of yeast for improving galactose utilization through integrated systems level analysis, Proc Natl Acad Sci U S A, 2011, 108(29), 12179–84. Brazma A, Hingamp P, Quackenbush J, et al. Minimum information about a microarray experiment (MIAME)-toward standards for microarray data, Nat Genet, 2001, 29(4), 365–71. Grigoriev I V, Cullen D, Goodwin SB, et al. Fueling the future with fungal genomics, Mycology, 2011, 2(3), 192–209. Regenberg B, Grotkjaer T, Winther O, et al. Growth-rate regulated genes have profound impact on interpretation of transcriptome profiling in Saccharomyces cerevisiae, Genome Biol, 2006, 7(11), R107. Richmond CS, Glasner JD, Mau R, Jin H, Blattner FR. Genome-wide expression profiling in Escherichia coli K-12, Nucleic Acids Res, 1999, 27(19), 3821–35. Selinger DW, Cheung KJ, Mei R, et al. RNA expression analysis using a 30 base pair resolution Escherichia coli genome array, Nat Biotechnol, 2000, 18(12), 1262–8. Andersen MR, Vongsangnak W, Panagiotou G, Salazar MP, Lehmann L, Nielsen J. A trispecies Aspergillus microarray: comparative transcriptomics of three Aspergillus species, Proc Natl Acad Sci U S A, 2008, 105(11), 4387–92. Bordel S, Agren R, Nielsen J. Sampling the solution space in genome-scale metabolic networks reveals transcriptional regulation in key enzymes, PLoS Comput Biol, 2010, 6(7), e1000859. Patil KR, Nielsen J. Uncovering transcriptional regulation of metabolism by using metabolic network topology, Proc Natl Acad Sci U S A, 2005, 102(8), 2685–9. Fernie AR, Stitt M. On the discordance of metabolomics with proteomics and transcriptomics: coping with increasing complexity in logic, chemistry, and network interactions scientific correspondence, Plant Physiol, 2012, 158(3), 1139–45. Trauger SA, Kalisak E, Kalisiak J, et al. Correlating the transcriptome, proteome, and metabolome in the environmental adaptation of a hyperthermophile, J Proteome Res, 2008, 7(3), 1027–35. Corbin RW, Paliy O, Yang F, et al. Toward a protein profile of Escherichia coli: comparison to its transcription profile, Proc Natl Acad Sci U S A, 2003, 100(16), 9232–7. Belle A, Tanay A, Bitincka L, Shamir R, O’Shea EK. Quantification of protein half-lives in the budding yeast proteome, Proc Natl Acad Sci U S A, 2006, 103(35), 13004–9.

56 | 2 Systems biology in biofuel

[42]

[43]

[44] [45] [46] [47]

[48] [49] [50] [51]

[52]

[53]

[54]

[55]

[56] [57] [58] [59] [60] [61]

Anderson KL, Roberts C, Disz T, et al. Characterization of the Staphylococcus aureus heat shock, cold shock, stringent, and SOS responses and their effects on log-phase mRNA turnover, J Bacteriol, 2006, 188(19), 6739–56. Yang Y, Liu B, Du X, et al. Complete genome sequence and transcriptomics analyses reveal pigment biosynthesis and regulatory mechanisms in an industrial strain, Monascus purpureus YY-1, Sci Rep, 2015, 5, 8331. Eichler J, Adams MWW. Posttranslational protein modification in Archaea, Microbiol Mol Biol Rev, 2005, 69(3), 393–425. Sibbald MJJB, Ziebandt AK, Engelmann S, et al. Mapping the pathways to staphylococcal pathogenesis by comparative secretomics, Microbiol Mol Biol Rev, 2006, 70(3), 755–88. Thanbichler M, Shapiro L. Getting organized–how bacterial cells move proteins and DNA, Nat Rev Microbiol, 2008, 6(1), 28–40. Verma S, Xiong Y, Mayer MU, Squier TC. Remodeling of the bacterial RNA polymerase supramolecular complex in response to environmental conditions, Biochemistry, 2007, 46(11), 3023–35. Picotti P, Bodenmiller B, Aebersold R. Proteomics meets the scientific method, Nat Methods, 2013, 10(1), 24–7. Domon B, Aebersold R. Mass spectrometry and protein analysis, Science, 2006, 312(5771), 212–7. Scherperel G, Reid GE. Emerging methods in proteomics: top-down protein characterization by multistage tandem mass spectrometry, Analyst, 2007, 132(6), 500–6. Mueller LN, Brusniak M-Y, Mani DR, Aebersold R. An assessment of software solutions for the analysis of mass spectrometry based quantitative proteomics data, J Proteome Res, 2008, 7(1), 51–61. Choe L, D’Ascenzo M, Relkin NR, et al. 8-plex quantitation of changes in cerebrospinal fluid protein expression in subjects undergoing intravenous immunoglobulin treatment for Alzheimer’s disease, Proteomics, 2007, 7(20), 3651–60. Ross PL, Huang YN, Marchese JN, et al. Multiplexed protein quantitation in Saccharomyces cerevisiae using amine-reactive isobaric tagging reagents, Mol Cell Proteomics, 2004, 3(12), 1154–69. Dong M, Yang LL, Williams K, et al. A “tagless” strategy for identification of stable protein complexes genome-wide by multidimensional orthogonal chromatographic separation and iTRAQ reagent tracking, J Proteome Res, 2008, 7(5), 1836–49. Huttlin EL, Hegeman AD, Harms AC, Sussman MR. Prediction of error associated with falsepositive rate determination for peptide identification in large-scale proteomics experiments using a combined reverse and forward peptide sequence database strategy, J Proteome Res, 2007, 6(1), 392–8. Elias JE, Gygi SP. Target-decoy search strategy for increased confidence in large-scale protein identifications by mass spectrometry, Nat Methods, 2007, 4(3), 207–14. Li X, Chen WN. Proteomics analysis of metabolically engineered yeast cells and mediumchained hydrocarbon biofuel precursors synthesis, AMB Express, 2014, 4(1), 61. Bradshaw RA, Burlingame AL, Carr S, Aebersold R. Reporting protein identification data: the next generation of guidelines, Mol Cell Proteomics, 2006, 5(5), 787–8. Wilkins MR, Appel RD, Van Eyk JE, et al. Guidelines for the next 10 years of proteomics, Proteomics, 2006, 6(1), 4–8. Oldiges M, Lütz S, Pflug S, Schroer K, Stein N, Wiendahl C. Metabolomics: current state and evolving methodologies and tools, Appl Microbiol Biotechnol, 2007, 76(3), 495–511. Soga T, Ohashi Y, Ueno Y, Naraoka H, Tomita M, Nishioka T. Quantitative metabolome analysis using capillary electrophoresis mass spectrometry, J Proteome Res, 2003, 2(5), 488–94.

References

[62]

[63]

[64]

[65] [66] [67]

[68] [69]

[70] [71] [72]

[73] [74] [75] [76] [77] [78] [79]

[80] [81] [82]

|

57

Bajad SU, Lu W, Kimball EH, Yuan J, Peterson C, Rabinowitz JD. Separation and quantitation of water soluble cellular metabolites by hydrophilic interaction chromatography-tandem mass spectrometry, J Chromatogr A, 2006, 1125(1), 76–88. Ewald JC, Heux S, Zamboni N. High-throughput quantitative metabolomics: workflow for cultivation, quenching, and analysis of yeast in a multiwell format, Anal Chem, 2009, 81(9), 3623–9. Fuhrer T, Heer D, Begemann B, Zamboni N. High-throughput, accurate mass metabolome profiling of cellular extracts by flow injection-time-of-flight mass spectrometry, Anal Chem, 2011, 83(18), 7074–80. Büscher JM, Czernik D, Ewald JC, Sauer U, Zamboni N. Cross-platform comparison of methods for quantitative metabolomics of primary metabolism, Anal Chem, 2009, 81(6), 2135–43. Giacomoni F, Le Corguillé G, Monsoor M, et al. Workflow4Metabolomics: a collaborative research infrastructure for computational metabolomics, Bioinformatics, 2015, 31(9), 1493–5. Villas-Bôas SG, Moxley JF, Akesson M, Stephanopoulos G, Nielsen J. High-throughput metabolic state analysis: the missing link in integrated functional genomics of yeasts, Biochem J, 2005, 388(Pt 2), 669–77. Schwarz D, Orf I, Kopka J, Hagemann M. Recent applications of metabolomics toward cyanobacteria, Metabolites, 2013, 3(1), 72–100. Canelas AB, Harrison N, Fazio A, et al. Integrated multilaboratory systems biology reveals differences in protein metabolism between two reference yeast strains, Nat Commun, 2010, 1, 145. Christen S, Sauer U. Intracellular characterization of aerobic glucose metabolism in seven yeast species by 13C flux analysis and metabolomics, FEMS Yeast Res, 2011, 11(3), 263–72. Zamboni N, Fendt S-M, Rühl M, Sauer U. (13)C-based metabolic flux analysis, Nat Protoc, 2009, 4, 878–92. Cortassa S, Caceres V, Bell LN, O’Rourke B, Paolocci N, Aon MA. From metabolomics to fluxomics: A computational procedure to translate metabolite profiles into metabolic fluxes, Biophys J, 2015, 108(1), 163–72. Dettmer K, Aronov PA, Hammock BD. Mass spectrometry-based metabolomics, Mass Spectrom Rev, 2007, 26(1), 51–78. Wiechert W. 13C metabolic flux analysis, Metab Eng, 2001, 3(3), 195–206. Niedenführ S, Wiechert W, Nöh K. How to measure metabolic fluxes: a taxonomic guide for (13)C fluxomics, Curr Opin Biotechnol, 2014, 34, 82–90. Winter G, Krömer JO. Fluxomics - connecting ’omics analysis and phenotypes, Environ Microbiol, 2013, 15(7), 1901–16. Bordbar A, Monk JM, King Z a, Palsson BO. Constraint-based models predict metabolic and associated cellular functions, Nat Rev Genet, 2014, 15, 107–20. Gerdes S, Edwards R, Kubal M, Fonstein M, Stevens R, Osterman A. Essential genes on metabolic maps, Curr Opin Biotechnol, 2006, 17(5), 448–56. Wecke T, Veith B, Ehrenreich A, Mascher T. Cell envelope stress response in Bacillus licheniformis: integrating comparative genomics, transcriptional profiling, and regulon mining to decipher a complex regulatory network, J Bacteriol, 2006, 188(21), 7500–11. Lerman J a, Hyduke DR, Latif H, et al. In silico method for modelling metabolism and gene product expression at genome scale, Nat Commun, 2012, 3(May), 929. Shannon PT, Reiss DJ, Bonneau R, Baliga NS. The Gaggle: an open-source software system for integrating bioinformatics software and data sources, BMC Bioinformatics, 2006, 7, 176. Mitra S, Klar B, Huson DH. Visual and statistical comparison of metagenomes, Bioinformatics, 2009, 25(15), 1849–55.

58 | 2 Systems biology in biofuel

[83] [84]

[85] [86] [87] [88]

[89] [90] [91] [92] [93] [94]

[95] [96] [97] [98]

[99]

[100]

[101] [102] [103]

Karp PD, Riley M, Saier M, Paulsen IT, Paley SM, Pellegrini-Toole A. The EcoCyc and MetaCyc databases, Nucleic Acids Res, 2000, 28(1), 56–9. Thomas A, Rahmanian S, Bordbar A, Palsson BØ, Jamshidi N. Network reconstruction of platelet metabolism identifies metabolic signature for aspirin resistance, Sci Rep, 2014, 4, 3925. Thiele I, Palsson BØ. A protocol for generating a high-quality genome-scale metabolic reconstruction, Nat Protoc, 2010, 5(1), 93–121. Feist AM, Palsson BØ. The growing scope of applications of genome-scale metabolic reconstructions using Escherichia coli, Nat Biotechnol, 2008, 26(6), 659–67. Price ND, Reed JL, Palsson BØ. Genome-scale models of microbial cells: evaluating the consequences of constraints, Nat Rev Microbiol, 2004, 2(11), 886–97. Garcia-Albornoz M, Thankaswamy-Kosalai S, Nilsson A, Väremo L, Nookaew I, Nielsen J. BioMet Toolbox 2.0: Genome-wide analysis of metabolism and omics data, Nucleic Acids Res, 2014, 42, W175–81. Schellenberger J, Que R, Fleming RMT, et al. Quantitative prediction of cellular metabolism with constraint-based models: the COBRA Toolbox v2.0, Nat Protoc, 2011, 6(9), 1290–307. Atsumi S, Hanai T, Liao JC. Non-fermentative pathways for synthesis of branched-chain higher alcohols as biofuels, Nature, 2008, 451(7174), 86–9. Tabatabaei M, Karimi K, Sárvári Horváth I, Kumar R. Recent trends in biodiesel production, Biofuel Res J, 2015, 2(3), 258–67. Lu X, Vora H, Khosla C. Overproduction of free fatty acids in E. coli: implications for biodiesel production, Metab Eng, 2008, 10(6), 333–9. Kalscheuer R, Stölting T, Steinbüchel A. Microdiesel: Escherichia coli engineered for fuel production, Microbiology, 2006, 152(Pt 9), 2529–36. Stöveken T, Kalscheuer R, Malkus U, Reichelt R, Steinbüchel A. The wax ester synthase/acyl coenzyme A:diacylglycerol acyltransferase from Acinetobacter sp. strain ADP1: characterization of a novel type of acyltransferase, J Bacteriol, 2005, 187(4), 1369–76. Liu T, Vora H, Khosla C. Quantitative analysis and engineering of fatty acid biosynthesis in E. coli, Metab Eng, 2010, 12(4), 378–86. Steen EJ, Kang Y, Bokinsky G, et al. Microbial production of fatty-acid-derived fuels and chemicals from plant biomass, Nature, 2010, 463(7280), 559–62. Duan Y, Zhu Z, Cai K, Tan X, Lu X. De novo biosynthesis of biodiesel by Escherichia coli in optimized fed-batch cultivation, PLoS One, 2011, 6(5), e20265. Teo WS, Ling H, Yu A-Q, Chang MW. Metabolic engineering of Saccharomyces cerevisiae for production of fatty acid short- and branched-chain alkyl esters biodiesel, Biotechnol Biofuels, 2015, 8, 177. Kalscheuer R, Luftmann H, Steinbüchel A. Synthesis of novel lipids in Saccharomyces cerevisiae by heterologous expression of an unspecific bacterial acyltransferase, Appl Environ Microbiol, 2004, 70(12), 7119–25. Yu KO, Jung J, Kim SW, Park CH, Han SO. Synthesis of FAEEs from glycerol in engineered Saccharomyces cerevisiae using endogenously produced ethanol by heterologous expression of an unspecific bacterial acyltransferase, Biotechnol Bioeng, 2012, 109(1), 110–5. Wang C, Yoon S-H, Jang H-J, et al. Metabolic engineering of Escherichia coli for α-farnesene production, Metab Eng, 2011, 13(6), 648–55. Peralta-Yahya PP, Ouellet M, Chan R, Mukhopadhyay A, Keasling JD, Lee TS. Identification and microbial production of a terpene-based advanced biofuel, Nat Commun, 2011, 2, 483. Peralta-Yahya PP, Keasling JD. Advanced biofuel production in microbes, Biotechnol J, 2010, 5(2), 147–62.

References

| 59

[104] Rude MA, Schirmer A. New microbial fuels: a biotech perspective, Curr Opin Microbiol, 2009, 12(3), 274–81. [105] Schirmer A, Rude MA, Li X, Popova E, del Cardayre SB. Microbial biosynthesis of alkanes, Science, 2010, 329(5991), 559–62. [106] Li N, Chang W-C, Warui DM, Booker SJ, Krebs C, Bollinger JM. Evidence for only oxygenative cleavage of aldehydes to alk(a/e)nes and formate by cyanobacterial aldehyde decarbonylases, Biochemistry, 2012, 51(40), 7908–16. [107] Li N, Nørgaard H, Warui DM, Booker SJ, Krebs C, Bollinger JM. Conversion of fatty aldehydes to alka(e)nes and formate by a cyanobacterial aldehyde decarbonylase: cryptic redox by an unusual dimetal oxygenase, J Am Chem Soc, 2011, 133(16), 6158–61. [108] Willis RM, Wahlen BD, Seefeldt LC, Barney BM. Characterization of a fatty acyl-CoA reductase from Marinobacter aquaeolei VT8: a bacterial enzyme catalyzing the reduction of fatty acylCoA to fatty alcohol, Biochemistry, 2011, 50(48), 10550–8. [109] Lennen RM, Pfleger BF. Microbial production of fatty acid-derived fuels and chemicals, Curr Opin Biotechnol, 2013, 24(6), 1044–53. [110] Rude MA, Baron TS, Brubaker S, Alibhai M, Del Cardayre SB, Schirmer A. Terminal olefin (1alkene) biosynthesis by a novel p450 fatty acid decarboxylase from Jeotgalicoccus species, Appl Environ Microbiol, 2011, 77(5), 1718–27. [111] de Jong B, Siewers V, Nielsen J. Systems biology of yeast: enabling technology for development of cell factories for production of advanced biofuels, Curr Opin Biotechnol, 2012, 23(4), 624–30. [112] Sukovich DJ, Seffernick JL, Richman JE, Hunt KA, Gralnick JA, Wackett LP. Structure, function, and insights into the biosynthesis of a head-to-head hydrocarbon in Shewanella oneidensis strain MR-1, Appl Environ Microbiol, 2010, 76(12), 3842–9. [113] Mendez-Perez D, Begemann MB, Pfleger BF. Modular synthase-encoding gene involved in α-olefin biosynthesis in Synechococcus sp. strain PCC 7002, Appl Environ Microbiol, 2011, 77(12), 4264–7. [114] Dash S, Mueller TJ, Venkataramanan KP, Papoutsakis ET, Maranas CD. Capturing the response of Clostridium acetobutylicum to chemical stressors using a regulated genome-scale metabolic model, Biotechnol Biofuels, 2014, 7(1), 144. [115] Visioli LJ, Enzweiler H, Kuhn RC, Schwaab M, Mazutti MA. Recent advances on biobutanol production, Sustain Chem Process, 2014, 2(1), 15. [116] Sillers R, Al-Hinai MA, Papoutsakis ET. Aldehyde-alcohol dehydrogenase and/or thiolase overexpression coupled with CoA transferase downregulation lead to higher alcohol titers and selectivity in Clostridium acetobutylicum fermentations, Biotechnol Bioeng, 2009, 102(1), 38–49. [117] Fontaine L, Meynial-Salles I, Girbal L, Yang X, Croux C, Soucaille P. Molecular characterization and transcriptional analysis of adhE2, the gene encoding the NADH-dependent aldehyde/alcohol dehydrogenase responsible for butanol production in alcohologenic cultures of Clostridium acetobutylicum ATCC 824, J Bacteriol, 2002, 184(3), 821–30. [118] Lütke-Eversloh T, Bahl H. Metabolic engineering of Clostridium acetobutylicum: recent advances to improve butanol production, Curr Opin Biotechnol, 2011, 22(5), 634–47. [119] Borden JR, Papoutsakis ET. Dynamics of genomic-library enrichment and identification of solvent tolerance genes for Clostridium acetobutylicum, Appl Environ Microbiol, 2007, 73(9), 3061–8. [120] Inui M, Suda M, Kimura S, et al. Expression of Clostridium acetobutylicum butanol synthetic genes in Escherichia coli, Appl Microbiol Biotechnol, 2008, 77(6), 1305–16. [121] Bond-Watts BB, Bellerose RJ, Chang MCY. Enzyme mechanism as a kinetic control element for designing synthetic biofuel pathways, Nat Chem Biol, 2011, 7(4), 222–7.

60 | 2 Systems biology in biofuel

[122] Nielsen J. Biofuels: chimeric synthetic pathways, Nat Chem Biol, 2011, 7(4), 195–6. [123] Shen CR, Lan EI, Dekishima Y, Baez A, Cho KM, Liao JC. Driving forces enable high-titer anaerobic 1-butanol synthesis in Escherichia coli, Appl Environ Microbiol, 2011, 77(9), 2905–15. [124] Knoshaug EP, Zhang M. Butanol tolerance in a selection of microorganisms, Appl Biochem Biotechnol, 2009, 153(1-3), 13–20. [125] Ezeji T, Milne C, Price ND, Blaschek HP. Achievements and perspectives to overcome the poor solvent resistance in acetone and butanol-producing microorganisms, Appl Microbiol Biotechnol, 2010, 85(6), 1697–712. [126] Krivoruchko A, Siewers V, Nielsen J. Opportunities for yeast metabolic engineering: Lessons from synthetic biology, Biotechnol J, 2011, 6(3), 262–76. [127] Fischer CR, Klein-Marcuschamer D, Stephanopoulos G. Selection and optimization of microbial hosts for biofuels production, Metab Eng, 2008, 10(6), 295–304. [128] Matsuda F, Furusawa C, Kondo T, Ishii J, Shimizu H, Kondo A. Engineering strategy of yeast metabolism for higher alcohol production, Microb Cell Fact, 2011, 10, 70. [129] Steen EJ, Chan R, Prasad N, et al. Metabolic engineering of Saccharomyces cerevisiae for the production of n-butanol, Microb Cell Fact, 2008, 7, 36. [130] Han M-J, Lee SY. The Escherichia coli proteome: past, present, and future prospects, Microbiol Mol Biol Rev, 2006, 70(2), 362–439. [131] Gasch AP, Werner-Washburne M. The genomics of yeast responses to environmental stress and starvation, Funct Integr Genomics, 2002, 2(4-5), 181–92. [132] Pham TK, Chong PK, Gan CS, Wright PC. Proteomic analysis of Saccharomyces cerevisiae under high gravity fermentation conditions, J Proteome Res, 2006, 5(12), 3411–9.

Nathalie Berezina

3 Production and application of chitin 3.1 Introduction For some years now, biopolymers have attracted great interest from both academia and industry. Some of them have been investigated for a long time, such as rubbers [1], the interest in others, such as starch, cellulose [2] or PHA [3], is mainly being driven by ecology concerns. Chitin is somehow apart from this mainstreaming interest in biopolymers. Indeed, as the second major biopolymer worldwide after cellulose, it is mainly produced as a byproduct in shellfish industry. Therefore its production was less a concern for its valorization through different applications and subsequent purifications and derivatizations [4, 5]. CH3 Glucosamine CH2OH H

O

H H OH

O

O

H H

H

OH H

N H O=C CH3

H

C=O CH2OH

N H H O

CH2OH

N–acetyl group

H

H O

H OH

O

O

H H

H

N H O=C CH3

n

Fig. 3.1: Chemical structure of chitin.

Chitin is a polysaccharide, more precisely an aminoglucopyranan, composed of Nacetylated glucosamine (GlcNAc) and glucosamine (GlcN) units (Fig. 3.1), linked by β(1,4) covalent bonds [6]. Due to this specific linkage, chitin exhibits an extremely robust structure towards chemical and biological aggression, indeed the β(1,4) chitin bond is similar to the one found in cellulose, contrary to starch, that is much more easily digested by enzymes of several (micro)organisms than cellulose and presents an α(1,4) covalent bond between its monomeric units. Moreover, the N-acetyl group attached to the major part of the glucosamine monomeric units of chitin confers to it extremely poor solubility properties, making chitin difficult to process and thus limiting its potential applications [4]. To circumvent this issue, the hydrolysis of the acetyl group, also called deacetylation, can be applied (Fig. 3.2). When GlcN units are predominant compared to GlcNAc units, the biopolymer is no longer designated as chitin, but as chitosan. Therefore the degree of acetylation (DA) (or the degree of deacetylation (DD)) is used to characterize the chitin/chitosan biopolymers as well as their degree of polymerization or source.

62 | 3 Production and application of chitin CH3 O HO O

O

OH

NH

O

O O

O

HO

NH

HO

O

Chitin

O

NH

OH

CH3

O

OH

CH3 Deacetylation

HO O

OH

NH2

O

O O

O O

HO

OH

CH3 NH

HO

O

Chitosan

O

NH2 OH

Fig. 3.2: Deacetylation of chitin to chitosan.

In this chapter the origin, different natural sources, subsequent issues related to the extraction and the purification of chitin, as well as its properties and applications are discussed.

3.2 Historical outline Chitin was most probably discovered by an English scientist A. Hachett, who reported in 1799 “a material particularly resistant to usual chemicals”, but as he did not push his investigations any further, the discovery of chitin is usually attributed to a French naturalist from Nancy, Henri Braconnot [7], who identified this biopolymer in 1811 in extracts of mushrooms, and thus gave it the name of “fungine” [8]. A few years later, in 1823, Auguste Odier found the same biopolymer in the insect’s exoskeleton, and named it “chitin” in reference to a Greek “chiton” meaning “tunic” [9]. The interrogation on the difference between cellulose, produced by plants, and chitin produced by arthropods was initiated by Payen in 1843 [9]. In the same year Lassaigne found the presence of nitrogen in chitin, when working with the exoskeleton of silkworm butterfly, Bombix morii [9]. Then, Ledderhose identified glucosamine and acetic acid as structural units of chitin in 1879, and Gilson confirmed glucosamine to be the repeated unit of chitin in 1894 [9]. The final chemical nature of chitin was elucidated by Purchase and Braun in 1946 [9].

3.3 Sources |

63

Chitosan was first obtained from chitin by C. Rouget [10], when boiling chitin in a concentrated alkali solution and noticing that the resulting compound was soluble in organic acids. Further, F. Hoppe-Seiler confirmed in 1894 that chitosan is the deacetylated form of chitin and thus gave it its actual name [10]. Another milestone in the discovery of chitin’s structure and arrangement was made by Bouligand. He discovered in 1965 that chitin adopts a stereotypic arrangement in arthropods [11]. Thus, three main types were found, α-, β- and γ-types (Fig. 3.3). The α-type exhibits an antiparallel disposal of chitin molecules, thus strengthening the intra- and inter-molecular hydrogen bonds. The α-type is the most robust type of chitin, it is the most resistant toward physical and chemical aggression and is also the one mainly found in nature. In contrast, the β-type chitin arrangement characterizes parallel chains’ aggregation, in this case N-acetyl groups play the role of spacers, allowing easier access to molecules of water, for hydration purposes and subsequent gel formations [11]. Finally, the γ-type is mainly composed by 2 parallel and 1 antiparallel layers, its strength and resistance are closer to the β-type, the N-acetyl groups playing the same role in both those arrangements. In addition, non-crystalline, transient states have also been reported in a fungal system [12].

α-chitin

β-chitin

γ-chitin

Fig. 3.3: Different chitin structures according to Bouligand.

Further, the macroscopic arrangement of chitin layers and protein scaffolds surrounding them on a cholesteric helix was studied [13], a twisted plywood structure was thus found in the lobster Homarus americanus [14] and in the sheep crab Loxorhynchus grandis [15], it has also been reported to be responsible for the iridescence of the scarab beetle [16].

3.3 Sources Chitin is present in the exoskeleton of arthropods [17], also in eukaryotic cells, such as those of fungi [18] and mushrooms [19]. It is also found in the iridophores (reflective material) in the epidermis and the eyes of certain arthropods and cephalopods [20]. One study [20] has even reported that the epidermal cuticle of a vertebrate, a fish named Paralipophrys trigloides, contains chitin; thus suggesting that chitin can also be produced by vertebrates. As chitin is present in so many different species, it would be, of course, very tempting to use chitin for evolution and taxonomy of these different species, however to the

64 | 3 Production and application of chitin

best of our knowledge those studies have still to be done. Also, a comparison between different properties, contents, etc. of chitin from different sources would be of great interest, but for now scientists could not distinguish any specific characteristics for chitin extracted from different sources (α-chitin being dominating in any shellfish, insects, etc.), also the content comparison faces the issue of chitin extraction and characterization, detailed below. Despite this very large diversity of chitin sources, until now, mainly chitin from the shellfish industry has been explored. It was thus shown that crustaceans mainly produce α-chitin, whereas cephalopods produce β-chitin, also the chitin content varies from 7 to 36 % in crustaceans and from 20 to 40 % in cephalopods, however we have to acknowledge that in this study for 10 species of crustaceans explored, only two representative of cephalopods were considered, therefore the conclusions concerning this latter class may be less accurate [21]. A few studies performed on insect chitin were mainly concerned with butterflies [22]. It was thus shown that the wings of the painted lady butterfly, Vanessa cardui Linnaeus, exhibits α-type chitin [22]. The insects exhibit a complex hierarchical structure, where each epidermal scale represents one color. These scales are morphologically homogeneous and adherent to the wings in rows, which run parallel to the anteriorposterior axis of the wing [22]. The chitin content is usually in the range of 5 to 40 % of dry mass of cuticle, thus honeybees were reported to have organic matrices with 23–32 % of chitin [17], mushrooms 8–16 % [17] and shellfish 5–40 % [21], and even up to 49 % were reported in squid pens [23]. The chitin content within the same species during the lifetime of the organism seems to remain stable, it was found to be around 3 % in Calanus helgolandicus [24]. Also, the determination of chitin content in its original source remains an issue. Indeed, no reliable analytic methods have been reported until now, infrared and diffraction analysis allowing only qualitative and non-quantitative approaches [25], therefore the only quantitative method mainly used up to now is the so called “alkaline extraction” method [25]. However, it strongly depends on the source and extraction procedure used, which may explain severe differences observed in the literature concerning some species, namely cuttlefish was found to have only 5.8 % of chitin by Hajji et al. [26] and up to 20 % by Rhazi et al. [21], whereas shrimp was found to have up to 37.2 % chitin by Hajji et al. [26] and only 22 % by Rhazi et al. [21].

3.4 Extraction and purification As we have seen from the previous paragraph, the extraction and purification of chitin is not only important for the recovery of the desired product but also to characterize the chitin content in different sources. Up to now two main approaches for chitin extraction have been studied: chemical and biological approaches.

3.4 Extraction and purification

|

65

3.4.1 Chemical extraction Chemical extraction basically consists of two steps: acidic treatment for mineral elimination and basic treatment for protein elimination [27, 28]. The classical procedure can consist of an acidic treatment with HCl 1 M for 2 to 24 hours, followed with a basic treatment with NaOH 1 M for another 24 to 48 hours. The temperature usually is kept at 60–90 °C to avoid as much as possible chitin degradation and preserve its structure. The different parameters of these treatments can be tuned in order to adapt to a specific source of chitin or to combine different steps [17, 29]. Thus, for example, the mineral content is especially high within shellfish sources of chitin whereas lipid content is much higher in insects. Therefore, the acidic treatment can be adjusted, especially through the solid-state approach, to combine the elimination of both these sources of contamination and to obtain highly crystalline chitosan [30]. Also the alkali concentration for the deproteination step is of specific interest, as at high temperature and concentration the reaction continues up to deacetylation and the obtained product is no longer chitin, but chitosan [31]. The utilization of acidic and basic conditions at high temperature may also damage the integrity of the chitin polymer, and oligosaccharides can be thus obtained. It can present advantages for some applications, shorter chains inducing better solubility, but also drawbacks of chain alteration [18]. Another issue consists of a very poor ability to analyze the obtained product, several attempts through infrared [32], diffraction [25], fluorescence [33] or NMR [21, 26] techniques were made, however they did not give sufficiently satisfying results [25, 28, 29], and only a combination of several of these approaches allows the confirmation of the structure and purity of the chitin. Another aspect of the purification of chitin is its color. Indeed, melanins and other sclerotins [12, 17] attached to the exoskeleton of arthropods or other chitin sources make it difficult to obtain a pure white chitin at the end of the process, therefore bleaching agents, such as hydrogen peroxide, are often used to finalize the chitin purification [34]. Other techniques, such as γ-radiation, were also tested to improve the extraction of chitin [35]. The main drawback of the utilization of chemical extraction remains however its energetic and environmental impact due to the extensive utilization of acidic and basic solutions requiring further neutralization and elimination [34].

3.4.2 Biological extraction The biological methods for the extraction of chitin have been developed recently [34]. They can use either purified enzymes [36, 37] or whole microorganisms [38].

66 | 3 Production and application of chitin

Although more environment friendly, these methods remain less efficient, one of the main reasons being the up to now poor understanding of the main covalent links between chitin and the surrounding proteins and melanins [12, 17]. Indeed, the main enzymes used for the purification of chitins are proteases, they cut protein bonds to give peptides and amino acids, however they cannot liberate chitin from catechols or even amino acids directly attached to it. In this case, it becomes even more difficult to avoid bleaching steps, thus reducing the scope of pure biological extraction and making it much more biochemical [34]. As neither chemical nor biological methods are completely satisfying for now, several academic teams and industrial companies are still working on their improvements. This extraction stage is, indeed, essential for chitin and chitosan to fulfill the huge potential those molecules can present.

3.5 Applications Chitin and chitosan have recently been reported to present several properties such as biocompatibility [39], biodegradation [40], scavenging of heavy metal [41] and of cholesterol or other fats [42], antimicrobial and antioxidative behaviors [43, 44], etc. However, we have to acknowledge that even if those biopolymers seem very promising, until now, the actual applications remain rather limited. This can be explained by different aspects: the difficulty of extraction and purification of the original chitin, the necessary transformation of chitin to chitosan followed by the derivatization of the latter for most of applications [45–47], etc. Also, some of the previously mentioned assertions appeared less obvious that they seemed at first glance, for example, it would be difficult for the same molecule to present antimicrobial and biodegradation properties, actually if the antimicrobial aspect of chitin and chitosan seems to be proven, their biodegradation appears much more doubtful [48]. Similarly, the fact that chitin is difficult to purify, namely from fats, does not necessarily mean that it makes it good candidate for lowering cholesterol in blood. For all the reasons mentioned above, we will focus our attention in this chapter mainly on biomedical, agricultural, materials and water purification applications of chitin and chitosan.

3.5.1 Biomedical applications The main biomedical applications of chitosan are in wound healing. This application combines two of the most interesting properties of chitosan: antimicrobial behavior and biocompatibility [4]. These products have been on the market since the

3.5 Applications

| 67

early 1990s, mainly in Asia and North America, but also in Europe, however to a lesser extent. These products are commercialized by companies such as BioSyntech (Canada), Hemcon (USA), Medovent (Germany), Marine Polymers Technologies (USA), Cytosial (France). Some other biomedical applications of chitosan were also studied, such as bone substitutes [4, 49], blood interactions [4, 50], drugs vectorization [4], implants [51], or anti-inflammatory [52, 53], antihypertensive [54] and anticancer [55] drugs; however most of these applications have not yet reached the actual market of biomedical products. Finally, chitosan was also shown to be active against cryptosporidiosis in goats, significantly reducing the excretion of oocysts of C. parvum and diarrhea, and enhancing the weight gain of young animals [56].

3.5.2 Agricultural applications Chitin and chitosan were reported to contribute to the protection of plants against pathogens. However, in this application, chitin and chitosan were described to play very different roles. Chitin was reported to contribute to the protection of seeds, mainly by allowing the growth of microbial pesticides, such as Trichoderma harzianum P1, which was reported to be active against foliar disease. The addition of chitin to the medium specifically contributed to the growth of T. harzianum and partially inhibited the growth of certain pathogens such as R. solani, thus playing a double role: enhancing the production of suitable microorganisms and lowering the invasion of unsuitable ones through actual inhibition and further demographic control [57]. Chitosan was described to act as an elicitor, indeed, several plants possess chitinolytic enzymes, which help them to defend themselves against pathogen aggressors, such as fungi. The introduction of chitosan in the growth medium stimulates the production of chitinolytic enzymes in plants, thus making them more resistant towards their natural aggressors [58, 59].

3.5.3 Materials applications For a long time chitosan as a material was used as a plastic for the production of antimicrobial films for the food industry, i.e. packaging protecting fresh vegetables, fruits or meat [60]. Several attempts to produce novel biofunctional materials were also made, such as grafting of ester derivatives of poly(ethylene glycol) (PEG) [61] or phosphomethylation [62, 63].

68 | 3 Production and application of chitin

More recently, some novel applications received greater interest, among them the utilization of chitosan as a catalyst support is of particular interest. Indeed, this application combines several up-to-date techniques, such as freeze drying [64] or utilization of supercritical CO2 (scCO2 ) [65] to increase the surface exchange capabilities and/or utilization of ionic liquids [66, 67]. It thus contributes to the implementation of green chemistry principles by minimizing the amount of required product – catalytic and not stoichiometric proportions, and utilization of renewable raw materials – second most abundant biopolymer worldwide.

3.5.4 Water purification The quality of water remains one of the huge issues humanity is currently facing. Access to pure water in some parts of the world remains difficult, thus generating malnutrition, diseases, and even military conflicts. Therefore the purification of water that was initially unsuitable for use or polluted is of crucial importance. To contribute to the wellbeing of a larger population the chosen water treatment techniques have to be technically and economically efficient. Different techniques are usually applied to water purification, such as scavenging, adsorption, flocculation or biological treatments. Chitosan can be applied for adsorption [68, 69] and flocculation [70] purposes, the fact that it is an environment friendly compound, very abundant and rather cheap makes it a solution of choice in certain cases. Thus chitosan was shown to be particularly efficient for the flocculation of cardboard-mill secondary biological wastewater [70], unfortunately the actual applications in industry remain rather rare, as concurrent flocculating agents are cheaper. Indeed, even if chitosan shows better properties, the traditional cheaper products are sufficient to fulfill current regulatory frameworks.

3.6 Outlook Chitin is a second most abundant biopolymer on Earth after cellulose. Present in several species, it is, until now, mainly obtained from the discards of the shellfish industry. Its extraction and purification as well as the reliability of available sources remain an issue and unfortunately impacts its widespread applications. Also, as chitin has a rather poor range of applications that can be exploited, mainly focused on the agricultural area, it has to be transformed by deacetylation to chitosan, and sometimes even further by derivatization of the latter. Despite extremely interesting properties, such as biocompatibility or antimicrobial behavior, the applications of chitosan in the biomedical area remain limited, mainly due to the extreme difficulty to access sufficient purity and source reliability of the biopolymer. The materials applications until now are rather limited, mainly

References

|

69

due to the cost of the product, which remains higher than that of petroleum based polymers with similar properties. The applications in the agricultural fields seem very promising but require more research and development to achieve significant results, therefore the main utilization of chitosan is now the purification of water, achieved by scavenging of heavy metal residues. Finally, for the widespread utilization of these truly very promising biopolymers, beside strong scientific developments, intellectual rigor is also mandatory, to promote only actual properties and to avoid attributing fashionable but not proven ones.

References [1] [2] [3]

[4] [5] [6] [7] [8] [9]

[10] [11] [12] [13] [14]

[15] [16] [17]

Martelli SM, Lin CSK, Sun Z, Berezina N, Fakhouri FM,-Mei LHI. Natural rubber blends with biopolymers, in Natural Rubber Materials: Volume 1: Blends and IPNs, RSC, 2013, 1, 370–93. Berezina N, Martelli SM. Bio-based polymers and materials In: Renewable resources for biorefineries, RSC Green Chemistry 2014, 27, 1–28. Berezina N, Martelli SM. Polyhydroxyalkanoates: structure, properties and sources. In: Polyhydroxyalkanoate (PHA) based blends, composites and nanocomposites, RSC Green Chemistry 2015, 30, 18–46. Khor H, Wan ACA. Chitin: fulfilling a biomaterials promise. Elsevier Insights, 2nd Ed., 2014. Archer M, Russel D. Crustacea processing waste management. Seafish, 2008, 1–23. Rebecca LJ, Susithra G, Sharmila S, Singh A. Optimization of physical parameters for chitinase production from Serratia marcescens, Res J Pharm Biol Chem Sci, 2013, 4, 1676–82. Wisniak J, Henri Braconnot. Revista CENIC. Ciencias Químicas, 2007, 38, 345–55 and references cited therein. Khoushab F, Yamabhai M. Chitin research revisited. Mar Drugs, 2010, 8, 1988–2012 and references cited therein. Discover polysaccharides: chitins and chitosans. [last accessed 2nd July 2015] http://polysac3db.cermav.cnrs.fr/discover_chitins_chitosans.html, and references cited therein. Teng D. From chitin to chitosan, in Chitosan-based hydrogels, Taylor & Francis Group, 2012, pp. 1–33 and references cited therein. Thomas S, Durand D, Chassenieux C, Jyotishkumar P, Handbook of biopolymer-based materials: from blends and composites to gels and complex networks, John Wiley & Sons, 2013. Merzendorfer H, Zimoch L. Chitin metabolism in insects: structure function and regulation of chitin synthases and chitinases, J. Exp. Biol., 2003, 206, 4393–412. Charvolin J, Sadoc JF. About collagen, a tribute to Yves Bouligand, Interface focus, 2012, 1–8. Raabe D, Romano P, Sachs C, Al-Sawalmih A, Brokmeier HG, Yi SB, Servos G, Hartwig HG. Discovery of a honeycomb structure in the twisted plywood patterns of fibrous biological nanocomposite tissue, J Crist Growth, 2005, 283, 1–7. Chen PY, Lin AYM, McKittrick J, Meyers MA. Structure and mechanical properties of crab exoskeletons, Acta Biomaerialia, 2008, 4, 587–96. Sharma V, Crneb M, Park JO, Srinivasarao J. Bouligand structures underlie circularly polarized iridescence ofscarab beetles: a closer view, Materials Today: Proc., 2014, 161–71. Nwe N, Furuike T, Tamura H. Chitin and chitosan from terrestrial organisms, in Chitin, chitosan, oligosaccharides and their derivatives, CRC Press, 2010, pp. 3–10.

70 | 3 Production and application of chitin

[18] Rinaudo M. Chitin and chitosan: Properties and applications, Prog. Polym. Sci., 2006, 31, 603–32. [19] Kurtzman RHJr. Mushrooms: sources for modern western medicine, Micalogia Aplicada Int., 2005, 17, 21–33. [20] Khousab F, Yamabhai M. Chitin research revisited, Mar. Drugs, 2010, 8, 1988–2012. [21] Rhazi M, Desbrieres J. Investigation of different natural sources of chitin: influence of the source and deacetylation process on the physicochemical characteristics of chitosan, Polymer Int., 2000, 49, 337–344. [22] Schiffman JD, Schauer CL. Solid state characterization of α-chitin from Vanessa cardui Linnaeus wings, Mat. Sci. Eng. C, 2009, 29, 1370–4. [23] Abdou ES, Nagy KSA, Elsabee MZ. Extraction and characterization of chitin and chitosan from local sources, Bioresour. Technol., 2008, 99, 1359–67. [24] Gervasi O, Jeuniaux C, Dauby P. Production de chitine par les crustacés du zooplancton de la baie de Calvie (Corse), IFREMER: Actes de colloques, 1988, 8, 33–8 in French. [25] Jeuniaux C, Voss-Foucart MF, Bussers JC. La production de chitine par les crustacés dans les écosystèmes marins, Aquat.Living Resour., 1993, 6, 331–41 in French. [26] Hajji S, Younes I, Ghorbel-Bellaaj O, Hajj R, Rinaudo M, Nasri M, Jellouli K. Structural differences between chitin and chitosan extracted from three different marine sources, Int. J. Biol. Macromol., 2014, 65, 298–306. [27] Shahidi F, Synowiecki J. Isolation and characterization of nutrients and value-added products from snow crab (Chinoecetes opilio) and shrimp (Pandalus borealis) processing discards, J. Agric. Food Chem., 1991, 39, 1527–32. [28] Rodde RH, Einbu A, Varum KM. Aseasonal study of the chemical composition and chitin quality of shrimp shells obtained from northern shrimp (Pandalus borealis), Carbohydr. Polym., 2008, 71, 388–93. [29] Lavall R, Assis OBG, Campana-Filho SP. Bioresour. Technol., 2007, 98, 2465–72. [30] Osorio-Madrazo A, David L, Trombotto S, Lucas JM, Peniche-Covas C, Domard A. Carbohydr. Polym., 2011, 83, 1730–9. [31] Plassard C, Mousan D, Salsac L. Dosage de la chitine sur des ectomycorhizes de pin maritime (Pinus pinaster) à Pisolithus tinctorius: évaluation de la masse mycélienne et de la mycorhization, Can. J. Bot., 1983, 61, 692–9. [32] Brugnerotto J, Lizardi J, Goycoole FM, Arguelles-Monal W, Desbrieres J, Rinaudo M. An infrared investigation in relation with chitin and chitosan characterization, Polymer, 2001, 42, 3569–80. [33] Rabasovic MD, Pantelic DV, Jelenkovic BM, Curcic SB, Rabasovic MS, Vrbica MD, Lazovic VM, Curcic NPM, Krmpot AJ. Nonlinear microscopy of chitin and chitinous structures: a case study of two cave-dwelling insects, J. Biomed. Optics, 2015, 20, 016010–10. [34] Vázquez JA, Rodríguez-Amado I, Montemayor MI, Fraguas J, del Pilar González M, Anxo Murado M. Chondroitin sulfate, hyaluronic acid and chitin/chitosan production using marine waste sources: Characteristics, applications and eco-friendly processes: A review, Mar. Drugs, 2013, 11, 747–74. [35] Mahlous M, Tahtat D, Benamer S, Nacer Khodja A. Gamma irradiation-aided chitin/chitosan extraction from prawn shells, Nucl. Instr. Meth. Phys. Res. B, 2007, 265, 414–7. [36] Duarte de Holanda H, Netto FM. Recovery of components from shrimp (Xiphopenaeus kroyeri) processing waste by enzymatic hydrolysis, J Food Sci., 2006, 71, 298–303. [37] Synowiecki J, Al-Khateeb NAAQ, The recovery of protein hydrolysate during enzymatic isolation of chitin from shrimp Crangon crangon processing discards, Food Chem., 2000, 68, 147–52. [38] Xu Y, Gallert C, Winter J. Chitin purification from shrimp wastes by microbial deproteination and decalcification, Environ. Biotechnol., 2008, 79, 687–97.

References

|

71

[39] Zhu A, Shen J. Biocativities of chitosan and its derivatives, in Chitosan-based hydrogels, Taylor & Francis, 2012, pp. 109–73. [40] Kumar MNVR. A review of chitin and chitosan applications, React. Funct. Polym., 2000, 46, 1–27. [41] Shepherd R, Reader S, Falshaw A. Chitosan functional properties, Glyconjug. J., 1997, 14, 535–42. [42] Koide SS. Chitin-chitosan: properties, benefits and risks, Nutr. Res., 1998, 18, 1091–101. [43] Kim KW, Thomas RL. Antioxidative activity of chitosans with varying molecular weights, Food Chem., 2007, 101, 308–13. [44] Park PJ, Koppula S, Kim SK. Antioxidative activity of chitosan, chitooligosaccharides and their derivatives, in Chitin, chitosan, oligosaccharides and their derivatives, CRC Press, 2010, pp. 241–50. [45] Mourya VK, Inambar NN. Chitosan-modifications and applications: opportunities galore, Reactive Funct. Polym., 2008, 68, 1013–51. [46] Tolaimate A, Desbrieres J, Rhazi M, Alagui A. Contribution to the preparation of chitins and chitosans with controlled physico-chemical properties, Polymer, 2003, 44, 7939–52. [47] Lai WF, Lin MCM. Chemical derivatization of chitosan for plasmid DNA delivery: present and future, in Chitin, chitosan, oligosaccharides and their derivatives, CRC Press, 2010, pp. 69–94. [48] Kirchman DL, White J. Hydrolysis and mineralization of chitin in the Delaware Estuary, Aquat. Microb. Ecol., 1999, 18, 187–96. [49] Muzzarelli RAA. Chitosan scaffolds for bone regeneration, in Chitin, chitosan, oligosaccharides and their derivatives, CRC Press, 2010, pp. 223–39. [50] Kim SK, Jung WK. Effects of chitin, chitosan and their derivatives on human hemostasis, in Chitin, chitosan, oligosaccharides and their derivatives, CRC Press, 2010, pp. 251–62. [51] Khor E, Lim LY. Implantable applications of chitin and chitosan, Biomaterials, 2003, 24, 2339–49. [52] Kim MM, Kim SK. Anti-inflammatory activity of chitin, chitosan, and their derivatives, in Chitin, chitosan, oligosaccharides and their derivatives, CRC Press, 2010, pp. 215–21. [53] Kim K, Ji HS. Effect of chitin sources on production of chitinase and chitosanase by Streptomyces griseus HUT 6037, Biotechnol. Bioprocess Eng., 2001, 6, 18–24. [54] Je JY, Ahn CB. Antihypertensive actions of chitosan and its derivatives, in Chitin, chitosan, oligosaccharides and their derivatives, CRC Press, 2010, pp. 263–70. [55] Ta HT, Dunstan DE, Dass CR. Anticancer activity and therapeutic applications of chitosan nanoparticles, in Chitin, chitosan, oligosaccharides and their derivatives, CRC Press, 2010, pp. 271–84. [56] Adjou K, Mammeri M, Grasset-Chevillot A, Marden JP, Auclair E, Mage C, Vallée I. Maitrise de la cryptosporidiose avec le chitosan: essi in vivo. Proceedings: Journées nationales DTV – Nantes 2015, pp. 377–80 in French. [57] Tronsmo A, Skaugrud O, Harman GE. Use of chitin and chitosan in biological control of plant diseases, in Chitin enzymology, Eur. Chitin Soc., 1993, pp. 265–70. [58] Kato J, Shimura Y, Sogitani M, Torii H, Nakajima K, Oishi K. Physiological role of chitinase and chitin-binding lectin in cucumber,in Chitin enzymology, Eur. Chitin Soc., 1993, pp. 257–64. [59] Lienart Y, Gautier C, Dubois-Dauphin R, Domard A. Tetramers of chitin (chitosan) as elicitors in Rubus protoplasts, in Chitin enzymology, Eur. Chitin Soc., 1993, pp. 271–6. [60] Nadarajah K, Prinyawtwatkui W, No HK, Sathivel S, Xu Z. Sorption behavior of crawfish chitosan films as affected by chitosan extraction processes and solvent types, J Food Sci., 2006, 71, 33–39.

72 | 3 Production and application of chitin

[61] Leduc F, Dez I, Desbrieres J, Picton L, Madec JP. Different ways for grafting ester derivatives of poly(ethylene glycol) onto chitosan: related characteristics and potential properties, Polymer, 2005, 46, 639–51. [62] Leduc F, Dez I, Madec JP. NMR study of the phosphonomethylation reaction on chitosan, Polymer, 2005, 46, 319–25. [63] Leduc F, Dez I, Gulea M, Madec JP, Jaffres PA. Synthesis of phosphorus-containing chitosan derivatives,in Phosphorus, sulfur, and silicon, Taylor & Francis, 2009, pp. 872–89. [64] Jouanin C, Vincent C, Dez I, Gaumont AC, Vincent T, Guibal E. Highly porous catalytic materials with Pd and ionic liquid supported on chitosan, J. Appl. Polym. Sci., 2013, DOI: 10.1002/app.38501. [65] Moucel R, Perrigaud K, Goupil JM, Madec PJ, Marinel S, Guibal E, Gaumont AC, Dez I. Importance of the conditioning of the chitosan support in a catalyst-containing ionic liquid phase immobilized on chitosan: the palladium-catalysed allylation reaction case, Adv. Synth. Catal., 2010, 352, 433–39. [66] Clousier N, Moucel R, Naik P, Madec PJ, Gaumont AC, Dez I. Catalytic materials based on catalyst containing ionic liquid phase supported on chitosan or alginate: importance of the support, C. R. Chimie, 2011, 14, 680–4. [67] Baudoux J, Perrigaud K, Madec PJ, Gaumont AC, Dez I. Developmetn of new SILP catalysts using chitosan as support, Green Chem., 2007, 9, 1346–51. [68] Crini G. Recent developments in polysaccharide-based materials used as adsorbents in wastewater treatment, Prog. Polym. Sci., 2005, 30, 38–70. [69] Crini G. Non-conventional low-cost adsorbents for dye removal: A review, K, 2005, DOI: 10.1016/j.biotech.2005.05.001. [70] Renault F, Sancey B, Charles J, Morin-Crini N, Badot PM, Winterton P, Crini G. Chitosan flocculation of cardboard-mill secondary biological wastewater, Chem. Eng. J., 2009, 155, 775–83.

Hui Li, Hu Zhu, Shiwei Sun, Zhimei Feng, Yajie Sun, and Wanlong Zhou

4 Biological production of welan gum 4.1 Sphingans: Occurrence and structure Since the genus Sphingomonas was identified in 1990, it has attracted much attention for several reasons. One reason is that these organisms have great potential to be applied in a wide range of biotechnological applications such as degradation of environmental pollutants, bioremediation and wastewater treatment. A more important reason is that they are an important microbial resource for biopolymer synthesis. The biopolymers they produce are bacterial exopolysaccharides (EPS) sharing similar backbone structures and are named sphingans. The known sphingans and their producing strains include heteropolysaccharide S-7 produced by Sphingomonas sp. ATCC 21423, gellan gum (S-60) produced by Sphingomonas elodea ATCC 31461, welan gum (S-130) produced by Sphingomonas sp. ATCC 31555, rhamsan gum (S-194) produced by Sphingomonas sp. ATCC 31961, diutan gum (S-657) produced by Sphingomonas sp. ATCC 53159, S-88 produced by Sphingomonas sp. ATCC 31554 and S-198 produced by Sphingomonas sp. ATCC 31853 [1]. Most of the sphingans have the conserved linear repeating tetrasaccharide backbone structure (glucose-glucuronic acid-glucose-X-, X may be L-rhamnose or L-mannose) except for S-7, in which the 2-deoxy glucuronic acid replaces the common glucuronic acid (Fig. 4.1). Variations in the nature and location of the side chains are observed in different sphingans, which lead to their unique rheological characteristics and different commercial applications. For example, in the native form of gellan, it has no sugar side chains but two acyl substituents (O-acetate and L-glycerate) linked to the C-2 or C-6 of the same D-glucose residue, respectively. The acyl side chains have a great effect on the gelling process and the hardness of the gel that it forms. Thus, chemical deacylation of the native gellan causes the gels it forms to change from a soft, elastic thermoreversible one to a harder, brittle one [2]. Commonly, commercially available gellan gum has three forms: Gelrite® (no acyl residue), Kelcogel® F (low acyl content) and Kelcogel® LT100 (high acyl content). Gelrite® is used as the substitute for agar in thermophiles cultivation and plant tissue culture media. It also has great potential to be applied in the biodegradation of gasoline, the gel-encapsulated bacteria transportation for bioaugmentation of contaminated aquifers and other environmental applications. Kelcogels are food-grade and can be used as gelling agent, stabilizer and suspending agent in food industries (such as dessert gels, sauces, puddings) and personal care industries (lotions, creams, and toothpastes) because of their advantages such as good thermal and acidic stability, adjustable gel elasticity and rigidity, high transparency and good taste-releasing ability [1]. Welan gum has acetyl and an

74 | 4 Biological production of welan gum R1 6 →3) β-D-Glcp(1→4)β-D-GlcA (1→4) β-D-Glcp (1→4) α-X(1→ 2 3 R2

R3

sphingan

R1

R2

R3

X

Gellan

L-Glycerate

O-Acetate

H

L-Rhap

S-88

O-Acetate

H

←1)-α-L-Rhap

L-Rhap or L-Manp

Diutan

O-Acetate

Rhamsan ←1)β-D-Glcp (6←1) β-D-Glcp Welan

H

O-Acetate ←1)-α-L-Rhap (4←1)-α-L-Rhap H

H

O-Acetate ←1)-α-L-Rhap or ←1)-α-L-Manp

L-Rhap L-Rhap L-Rhap

(a) β-D-Glcp (1→6)β-D-Glcp 1 ↑ 6 →3) β-D-Glcp (1→4)β-D-2dGlcA (1→4) β-D-Glcp (1→4) α-L-Rhap (1→ 2 3 H

H

(b) Fig. 4.1: Repeat tetrasaccharide backbone structures of typical sphingans. (a) The sphingans with the conserved repeat tetrasaccharide backbone structure glucose-glucuronic acid-glucose-X- (X may be L-rhamnose or L-mannose). Glcp, GlcA, Rhap and Manp represent glucose, glucuronic acid, rhamnose and mannose, respectively. One special point is that the linkage of the acetyl residue in S-88 is unknown in the table. (b) Structure of S-7 in which the 2-deoxy glucuronic acid (2dGlcA) replaces the glucuronic acid in the backbone.

L-rhamnosyl or L-mannosyl as a side group. It does not gel but it presents excellent rheological properties at high temperatures and can be applied in petroleum, food, concrete and many other industries. Rhamsan has two D-glucosyl side chains and it can be used as the suspending agent for pesticides and fertilizers due to its tolerance to high concentrations of ammonium phosphate and salt solutions. Diutan has as a two rhamnose side chain while S-88 has only one rhamnose side chain. These sphingans are usually used as thickening agents (Tab. 4.1). Among these biopolymers, more attention is being paid to gellan because of its novel properties and industrial applications. However, other sphingan group biopolymers like welan gum also have attracted much attention and it is necessary to explore their potency. Welan gum is an anionic polysaccharide and has polyelectrolyte properties due to the presence of D-glucuronic acid in its chemical structure. Welan gum has great potential to be applied in many industries. For example, compared with the commonlyused water-soluble polymer xanthan gum, welan gum showed higher temperature stability (stable up to 150 °C which is 20 °C higher than that of xanthan gum) and oil

4.2 Welan gum: structure and properties |

75

Tab. 4.1: The properties and applications of different sphingans. Sphingan

Properties

Possible Applications

Gellan

It produces a thermoreversible gel when heated and cooled in the presence of cations.

Food, medical, agar substitute

Welan

Non-gelforming, but it produces high viscous solutions thermally stable up to 150 °C, relatively compatible with high concentrations of divalent ions and resistant to pH from 2 to 12

Oil field, cement, food, ink

S-88

Rheological properties such as thermal stability up to 150 °C are similar to welan

Printing pastes, oil drilling fluid

Diutan

The most thermally stable among all the known sphingans

Oil field, cement

Rhamsan

Similar to those of welan. It is more viscous at low concentration and low shear, and has a greater resistance to shear forces, but it loses its viscosity more than welan at temperatures above 100 °C. It tolerates high concentrations of phosphates and sodium chloride

Food, paints, coatings, plastic surgery, fertilizers, and pesticides

S-7

High viscosity in aqueous solutions, excellent suspending ability exceeding that of most other polymers, but it is sensitive to acid hydrolysis

Oil drilling fluid, a dripless water-based paint, or anti-icing composition for airplane surfaces

displacement efficiency, which indicates that it might be a good oil-displacing agent in oil-well drilling. High salt and pH tolerance makes welan gum an ideal viscosifer for cement-related applications where it is desirable to prevent phase separation. This chapter elaborates welan gum structure, production, biosynthetic pathway, and its potential applications. Engineering approaches for improvement of sphingan production are also discussed.

4.2 Welan gum: structure and properties The structure of welan gum has been identified in the following procedure: It is firstly subject to partial acid hydrolysis and base-catalyzed β-elimination to yield a series of oligosaccharides. These oligosaccharides are then converted into their alkylated alditol derivatives and analyzed by reverse-phase HPLC, fast atomic bombardment mass spectrometry (FABMS), proton nuclear magnetic resonance (1 H NMR) spectroscopy and gas liquid chromatography (GLC). The results show that the backbone

76 | 4 Biological production of welan gum

structure of welan gum is identical to the gellan family and composed of tetrasaccharide repeating units of D-glucose, D-glucuronic acid, D-glucose, and L-rhamnose. However, unlike gellan, it has an additional L-rhamnosyl or L-mannosyl side chain substituted on C3 of the first glucose residue of the repeating units and the ratio of occurrence between rhamnose and mannose is about 2 : 1 [3]. The molar ratio of L-mannose, L-rhamnose, D-glucose and D-glucuronic acid is 1.0 : 4.5 : 3.1 : 2.3 and the content of D-glucuronic acid is about 11.6–14.9 %. Besides, at least 85 % of the repeat units also have an acetyl substituent at C2 of the second glucose [4]. The molecular weight (MW) of welan gum is approximately 1.0 × 106 g/mol [5]. The characteristic chemical structure of welan gum accounts for its unique physiochemical properties and technological applications. Different from gellan gum, welan gum forms solutions with high viscosity instead of gels. The welan gum aqueous solution has the typical behavior of a pseudoplastic fluid because its viscosity decreases gradually as the shear rate increases and recovers when the share rate decreases [6]. Welan gum in aqueous media shows excellent rheological properties at low shear rate. It has good temperature endurance and no viscosity loss occurs upon autoclaving at 121 °C for 15–20 min [5]. Welan gum has shown to be chemically stable at 150 °C, which is much higher than that of xanthan gum (130 °C). Its high temperature stability might be related to its double-helical structure stabilized by side chain-main chain interactions. Besides, its aqueous solution also has good stability over a broad range of pH (2–12) and various salt concentrations (especially Ca2+ ). The good stability of welan gum makes it highly interesting for many applications such as oil-well drilling and the cement industry.

4.3 Production of welan gum 4.3.1 Producing strains At present, the known welan gum producing strains are Sphingomonas sp. (previously were divided into Alcaligenes), one group of typical Gram-negative, rod-shaped, chemoheterotrophic and aerobic bacteria whose cell envelopes contain glycosphingolipids instead of lipopolysaccharide and they usually form yellow-pigmented colonies. One commonly-used welan gum producing strain is Sphingomonas sp. ATCC 31555, (also known as Alcaligenes sp. ATCC 31555) [5]). Recently, more welan gum producing strains including Alcaligenes sp. NX-1 (CGMCC No.2428 ) [7], Alcaligenes sp. NX-3 [8] and Sphingomonas sp. TP-5 (CGMCC No.3097) [9] have been isolated from soil, wastewater and other environments. Furthermore, to enhance the production of welan gum, some mutants have been obtained by mutation breeding of the natural strains. One high welan gum producing mutant strain of Alcaligenes sp. NX-3, Alcaligenes sp. NX-3-1 has been developed using low-energy ion beam implantation technique and its welan gum yield reached 25.0 g/l after 66 h of cultivation in a 7.5 l

4.3 Production of welan gum

| 77

bioreactor, which was 34.4 % higher than that of the wild-type strain [10]. Another high-temperature-tolerant-producing mutant, Sphingomonas sp. HT-1 was obtained by atmospheric and room temperature plasma-induced mutation breeding of the strain Sphingomonas sp. CGMCC1737. Its optimal temperature is 37 °C which is much higher than that of the wild strain (the optimal temperature is 30°C). The mutant shows an improved production and viscosity of welan gum. Its maximum concentration of welan gum is 26.8 ± 0.34 g/l at 37 °C while that of the wild-type is 21.4 ± 0.35 g/l at 30 °C. Compared with the original strain, the welan gum viscosities of 1 % aqueous and 1 % KCl produced by the mutant increased by 41.7 % and 11.6 %, respectively [11].

4.3.2 Producing conditions 4.3.2.1 Fermentation media compositions Welan gum is the microbial fermentation product. The composition of the fermentation medium is very important for the growth of microorganism and the production of welan gum. Many methods, such as single factor experiment, orthogonal test, Plackett–Burman design and central composite design, have been applied to optimize the composition of the fermentation medium. Usually, carbon sources, nitrogen sources and mineral salts in the fermentation medium used for welan gum fermentation have a great impact on its production.

4.3.2.1.1 Carbon sources As one necessary substance for microorganism growth, a suitable carbon source will also improve the production of welan gum. Like other EPS such as gellan gum, rhamsan gum and xanthan gum, the microorganism can utilize commonly-used carbon sources including glucose, sucrose, starch, maltose or lactose to synthesize welan gum. Li et al. [8] used six different carbon sources including glucose, sucrose, maltose, corn starch, lactose, glycerol, citric acid and fumaric acid in welan gum fermentation to search for the optimal carbon sources. They found that when corn starch served as the carbon source, the biomass, the broth viscosity and the yield of welan gum were highest, followed by sucrose and glucose. Utilization of lactose by Alcaligenes facalis NX-3 was poor, and the production of welan gum was very low. The strain in the medium with glycerol, citric acid or fumaric acid was difficult to grow. Zhao et al. [12] observed that sucrose was the optimal carbon source among glucose, sucrose, starch and cassava starch hydrolysate. To reduce the production cost, using a cheap carbon source to obtain a high yield of welan gum is very important for its large-scale industrial production. So, some cheap carbon sources like waste from the dairy industry (whey), sugar industry (molasses and sugar beet) are attractive choices for welan gum production.

78 | 4 Biological production of welan gum

4.3.2.1.2 Nitrogen sources Nitrogen sources provide nitrogen for microbial growth. Nitrogen is one important element in protein and nucleic acids and accounts for 12.15 % of the cell dry weight. Like carbon sources, nitrogen sources are the main nutrients for microorganisms and have a great influence on microbial growth and synthesis of EPS. Among the seven nitrogen sources (peptone, yeast extract, defatted soybean powder, defatted peanut powder, cottonseed cake flour, ammonium nitrate and ammonium sulfate), the highest yield of welan gum was obtained by cottonseed cake flour; highest biomass was obtained by yeast extract; inorganic nitrogen sources (NH4 )2 SO4 and NH4 NO4 had a poor effect on the biomass and welan gum production [8]. Zhao et al. [12] found that the optimal nitrogen source was yeast extract and the poor nitrogen sources were inorganic nitrogen sources when five different nitrogen sources (peptone, corn steep liquor, yeast extract, ammonium nitrate, ammonium sulfate and urea) were used to search for the optimal nitrogen source.

4.3.2.1.3 Metal ions The concentration of metal ions in the medium has an important impact on the production of microbial polysaccharides. Metal ions play an important role in maintaining the conformation of the active sites and activity of enzymes that participate in the biosynthetic pathway of polysaccharides. Usually, metal ions such as Ca2+ , Mg2+ , Mn2+ , and Fe3+ may affect polysaccharides production and be served as substrates or cofactor in the polysaccharide synthetic process. In another respect, metal ions have a significant effect on the structure and properties of the synthesized biopolymer. It has been proved that the presence of Ca2+ affects the structural stability of welan gum by enhancing the interactions between double helices of the welan gum molecule [4].

4.3.2.1.4 Additives In terms of the microbial polysaccharides (cellulase, gellan gum and xanthan gum etc.), additives including precursors, enzymes surfactants, polar organic solvents and other substances can enhance their production by fermentation via increasing cell membrane permeability and improving the mass transfer environment. In the process of welan gum fermentation, addition of precursors like UDP-glucose, UDPglucuronic acid, dTDP-rhamnose and other substances that are necessary for welan gum biosynthetic pathway might enhance its yield. Li et al. [13] studied the effects of supplementation of two precursors, glucose-6-phosphate or fructose-6-phosphate or their mixture on welan gum production. The results suggested that the production of welan gum was improved by 18 % when their mixture was added. The effects of 11 additives (cetyltriethylammonium bromide, sodium dodecyl sulfate, Triton X-100, Span-20, Span-40, Span-80, Tween-20, Tween-40, Tween-80, DMSO and glycerol) on the biosynthesis of welan gum by Alcaligenes sp. CGMCC2428 were investigated.

4.3 Production of welan gum |

79

Tween-40, DMSO and glycerol were the most effective additives and the highest welan gum concentration (22.50 ± 0.65 g/l) was obtained with Tween-40, the second (21.15 ± 0.70 g/l) was obtained with glycerol and the third (20.51 ± 0.61 g/l) was obtained with DMSO and they were higher than that (19.05 ± 0.72 g/l) without additives [14].

4.3.2.2 Fermentation conditions Besides fermentation media compositions, the fermentation conditions such as temperature, pH, dissolved oxygen (DO) and shear stress also exhibit great influence on welan gum production.

4.3.2.2.1 Temperature and pH Temperature is very important for enzyme activity. The optimum temperature during the fermentation process will be beneficial for welan gum synthesis. The initial pH of the fermentation medium determines the dissociated status of the nutrients, restricts the rate of nutrients utilized by microbial cells, and directly affects intracellular enzyme activity. For the fermentation of welan gum, the optimum temperature is between 28–35 °C and pH ranges from 6.5–7.5. Li et al. [15] investigated the effects of different pH values (ranging from 5.5 to 7.0) on welan gum production in batch fermentation and the results showed that the dry cell weight and welan gum concentration increased as pH increased and reached maximum when pH was maintained at 7.0. So, the culture pH was controlled at 7.0 and the welan concentration (25.1 ± 0.65 g/l) and productivity (0.42 ± 0.003 g/l/h) was significantly improved compared with those (18.5 ± 0.72 g/l and 0.28 ± 0.002 g/l/h) obtained in native pH process. The metabolic flux distribution indicated that this improvement might result from the increased metabolic flux of α-glucose-1-phosphate (α-G1P) to welan in controlled pH process (12.3 %) compared with that (9.6 %) in the native pH process. Scale-up fermentation under controlled pH was performed in 300-L fermentor and the welan gum concentration reached 27.5 ± 0.97 g/l.

4.3.2.2.2 Dissolved oxygen (DO) and shear stress The synthesis of EPS (such as xanthan gum, gellan gum and hyaluronic acid) and its MW is affected by dissolved oxygen (DO) and shear stress of the fermentation process. In the fermentation process, the broth viscosity will increase due to the accumulation of EPS, and leads to a significant decrease in the oxygen mass transfer rate which is essential for the growth of microorganisms. The effects of DO on the synthesis of EPS might be different and complicated. For example, Banik and Santhiagu observed that the DO tension above 20 % had no effect on cell growth but gellan yield increased as the DO tension increased and reached 23 g/l when the DO tension level is 100 % [16]. Interestingly, Giavasis et al. [17] offered a different opinion on the effect of oxygen

80 | 4 Biological production of welan gum

supply upon the synthesis and MW of gellan gum. They found that the production of gellan gum did not increase as agitation increased, while the highest MW and concentration of gellan gum was obtained at moderate agitation. To ensure efficient welan gum production with high yield and high viscosity, Li et al. [18] established a proper oxygen supply strategy based on agitation speed control in batch fermentation of welan gum by Alcaligenes sp. CGMCC2428. They observed that the production of welan gum was affected by agitation speeds and the relatively higher maximum welan gum concentrations were obtained at the agitation speeds of 600 and 800 rpm. The kinetics analysis of the fermentation process at the two different agitation speeds suggested that cell growth and glucose consumption were faster at the beginning of fermentation when the agitation speed was at 800 rpm and as the fermentation proceeded, the agitation speed of 600 rpm was better for welan gum formation. So a two-stage agitation speed control strategy was proposed as follows: the agitation speed was controlled at 800 rpm for fast cell growth and glucose consumption in 22 h and the agitation was reduced to 600 rpm step-wise to maintain a stable dissolved oxygen level and obtain high welan gum production (26.3 ± 0.89 g/l) and yield (0.53 ± 0.003 g/g). Then the effects of DO and shear stress on the synthesis and MW of welan gum produced by Alcaligenes sp. CGMCC2428 were studied. The results showed that a high shear stress (1000 rpm) enhanced cell growth, but decreased MW and welan gum concentration. The maximum welan gum concentration (22.8 ± 0.61 g/l) and highest MW (9.01 ± 0.12 × 105 Da) was obtained at 800 rpm and 600 rpm, respectively. A moderate DO concentration (20 %) was suitable for both higher welan gum synthesis and its MW. The metabolic flux distribution of the strain at various DO concentrations was also analyzed to further investigate the effects of DO concentration. The outcomes demonstrated that the flux of α-G1P to welan gum was enhanced at 20 % of DO concentration, which might result in increased welan gum concentration and MW [19].

4.3.3 Recovery and purification of welan gum Besides welan gum, many impurities including cells, carbohydrates residuals and inorganic salts also exist in the fermentation broth and influence the quality of the welan gum products. Furthermore, the high viscosity of fermentation broth makes it difficult to extract the pure product and increases the extraction cost. So, it is of great importance to find a suitable and low cost method for welan gum recovery. The most common method for recovery of polysaccharide from fermentation broth in industry is the precipitation method with alcohol such as methanol, ethanol and isopropanol [20]. The polysaccharide can be easily recovered by centrifugation after it is precipitated by ice alcohol. Then the product can be dried at 50–60 °C in a forced-air tray drier or in a hot air oven or under partial vacuum. One advantage of this method is that alcohol is very easy to remove by evaporation and makes the purification of welan gum very simple. Polyethylene glycol (PEG) has also been used to precipitate

4.4 Biosynthetic pathway of welan gum

| 81

polysaccharide from cell free fermentation broth after the broth was treated with sulfuric acid and sodium chloride [21]. Usually, the pH of fermentation broth is adjusted before precipitation with alcohol in the preparation of rapidly hydrating welan gum. Many methods can be used in the purification of welan gum. One method is as follows: the crude gum is dissolved in NaCl and precipitated with isopropanol. Then the precipitate is washed with water and isopropanol several times to remove impurities and finally dried under vacuum. The second method is alkaline treatment by NaOH. The diluted sample is dissolved in the aqueous solution of NaOH and then neutralized with HCl. The neutralized sample is centrifuged and filtered on a membrane with pore size below 0.2 µm. This method resulted in a decrease in nitrogen and aggregate contents. The third method is using centrifugation and filtration to remove insoluble material and a proportion of high soluble mass material, and dialysis to remove low molar mass impurities. Other promising techniques such as size-exclusion chromatography and ion-exchange chromatography still need to be developed for the purification of welan gum [22].

4.4 Biosynthetic pathway of welan gum

Phosphoglucomutase

we lB rm lA rm lC rm lB rm lD urf 31 . urf 4 31 urf 34

atr B

we lF we lD we lC we lE we lM we lN atr D

we lG we lS we lR we lQ we ll we lk we lL we lJ

we lA

pG

ug dG

ug

pg

mG

In general, the biosynthetic scheme of sphingans such as gellan gum, S-88, S-7 and diutan gum follows the Wzx/Wzy-dependent pathway and can be divided into three sequential steps: (i) intracellular synthesis of nucleotide-sugar precursors, (ii) assembly of the tetrasaccharide repeating units attached at an undecaprenyl phosphate carrier by several glycosyltransferases, and (iii) polymerization of the assembled repeating units and export of the polysaccharide. Genes for biosynthesis of sphingan S-88, S-7, gellan gum, diutan gum have been elucidated. As observed for many other EPS, main genes necessary for their biosynthesis except those necessary for nucleotide sugar precursors synthesis (such as UDP-D-glucose and UDP-D-glucuronic acidare, but not dTDP-rhamnose) are clustered in the genome. The genome of the welan gum producing strain, Sphingomonas sp. ATCC 31555, has been sequenced and ten coding sequences (CDs) responsible for welan gum synthesis and 55 CDs asso-

Two component regulatory protein Lyase

UDP-Glucose pyrophosphorylase Polymerization/Export

UDP-Glucose dehydrogenase Translocase

Glycosyltransterase

Unknown function

ABC-transporter

dTDP-L-rhamnose biosynthesis

Fig. 4.2: Organization of the welan gum biosynthetic wel clusters from Sphingomonas sp. ATCC 31555. Arrows represent ORFs, and gene designations are indicated above the arrows. Known or putative gene functions are indicated.

82 | 4 Biological production of welan gum

ciated with monosaccharide metabolism have been annotated [23]. By comparing the different clusters for sphingan biosynthesis and available sequence data in the public databases, Schmid et al. [24] presented a wel cluster responsible for welan gum biosynthesis based on the genome sequence of Sphingomonas sp. ATCC 31555. We also analyzed the same genome and elucidated a similar wel cluster. The organization of this wel cluster is shown in Fig. 4.2 and Tab. 4.2 according to the locus of genes on the genome and their putative functions. Based on the cluster, we will discuss the proposed biosynthetic pathway of welan gum (Fig. 4.3) in this section. Tab. 4.2: Comparative analysis of putative proteins in welan gum production and corresponding proteins for other known sphingan synthesis. Putative proteins

Predicted function

Proteins in synthesis of dTDP-rhamnose RmlA glucose-1-phosphate thymidilyltransferase RmlB dTDP-D-glucose-4,6-dehydratase RmlC dTDP-4-dehydrorhamnose-3,5-epimerase RmlD dTDP-4-dehydrorhamnose-reductase

Identity (%) gel sps dps 95 97 93 88

99 99 96 85

97 99 95 87

Assembly of the tetrasaccharide repeating unit WelB glucosylisoprenylphosphate transferase WelK glucosyl-α-pyrophosphorylpolyprenol (PPL)-1,4-glucuronosyltransferase WelL glucosyl-glucuronosyl-transferase WelQ Rhamnosyl transferase

84 86

85 88

83 87

71 65

75 73

75 64

Polymerization/Export WelR Putative polysaccharide lyase WelG Putative polysaccharide polymerase WelS Putative flippase or repeat unit transporter WelI Polysaccharide secretion/attachment WelJ Polysaccharide secretion (ATPase) WelD Polysaccharide export protein WelC Export protein (chain length determinant) WelE Export protein (tyrosine kinase) WelM Polysaccharide secretion/attachment WelN Polysaccharide secretion/attachment atrD Secretion protein atrB ABC transporter

52 44 61 76 75 68 79 75 78 72 80 89

62 57 76 73 76 73 80 76 80 72 81 88

60 48 22 72 75 72 78 78 77 72 78 88

Regulation WelA Sensor kinase and response regulatory proteins

85

—

—

Unknown functions Urf31.4 Unknown Urf31 Putative glycosyltransferase Urf34 Membrane transport protein WelF Unknown, essential for polysaccharide synthesis

— — 89 57

— 80 88 58

75 82 88 55

4.4 Biosynthetic pathway of welan gum |

83

Glc

ATP GK ADP

PGI α-G6P

PgmG

β-F6P PMI

RmlA α-M6P

dTDP-Glc

α-G1P

UgpG

NADPH

RmlB

NADP+

RmlD

PPi

RmlC

α-M1P

PPi

UgdG

GTP

UTP

dTDP-Rha

UDP-GlcA UDP-Glc NAD(P)+ NAD(P)H, H+

GDP-Man

P P

P P

P P

UDP

UDP

UDP-Glc

P P

WelQ dTDP-Rha

UDP

dTDP

P P

urf31?

Ac-CoA

GDP-Man

C55-isoprenyl

UDP-GlcA

WelL

C55-isoprenyl

UDP-Glc

WelK

C55-isoprenyl

WelB

C55-isoprenyl

C55-isoprenyl

C55-isoprenyl

Step I synthesis of the nucleotide-sugar precursors in cytoplasm

P P GDP

Step II assembly of the tetrasaccharide repeating unit

OM WelD

C55-isoprenyl

P P

IM

WelE

WelC

WelG

P P

WelS

C55-isoprenyl

periplasm

cytoplasm Step III polymerization and export

Fig. 4.3: Proposed biosynthetic pathway for welan gum with mannose as the side chain. The process can be divided into three sequential steps: (i) intracellular synthesis of nucleotide-sugar precursors via PgmG, UdpG, UgpG, etc., (ii) assembly of the tetrasaccharide repeating units occurring at the lipid linkers anchored in the inner membrane by several glycosyltransferases and (iii) polymerization of the assembled repeating units and export of the polysaccharide. Abbreviations: Glc: Glucose; α-G6P: α-glucose-6-phosphate; β-F6P: β-fructose-6-phosphate; α-G1P: α-glucose-1-phosphate; α-M6P: α-mannose-6-phosphate; α-M1P: α-mannose-1-phosphate; UDP-Glc: UDP-glucose; UDP-GlcA: UDP-glucuronic acid; dTDP-Glc: dTDP-glucose; dTDP-Rha: dTDP-rhamnose; GDP-Man: GDP-mannose; Ac-CoA: acetyl coenzyme A; OM: outer membrane; IM: inner membrane.

84 | 4 Biological production of welan gum

4.4.1 Synthesis of the nucleotide-sugar precursors Like other sphingans, the biosynthetic pathway of welan gum starts with the formation of the nucleotide-sugar precursors. Li et al. [13] proposed the biosynthetic pathway of nucleotide-sugar precursors essential for welan gum production in strain Alcaligenes sp. CGMCC2428 by detecting the presence of several intermediates and key enzymes, investigating the effects of addition of precursors to the culture medium and correlating the activities of key enzymes with the yields of welan gum. The synthetic pathway is divided into cell metabolism and welan gum synthesis. After the uptake step of the extracellular glucose via an active transport system, glucose is metabolized by the cell through two possible ways. One is oxidative periplasmic pathway requiring GDH enzyme that uses pyrroloquinoline quinone as a cofactor to produce gluconate. Then gluconate is phosphorylated by an ATP-dependent gluconokinase reaction which produces 6-phosphogluconate (6-P-G). The other is phosphorylation to produce α-glucose-6-phosphate (α-G6P) by glucokinase. The α-G6P is further converted into β-fructose-6-phosphate (β-F6P) and α-G1P by phosphoglucoisomerase (PGI; EC 5.3.1.9) and phosphoglucomutase (PgmG; EC 5.4.2.2), respectively. Then two routes derive from α-G1P. One produces UDP-D-glucose and UDP-D-glucuronic acid catalyzed by pyrophosphorylase (UgpG; EC 2.7.7.9) and UDP-glucose dehydrogenase (UgdG; EC 1.1.1.22), respectively. The other route forms dTDP-L-rhamnose by four enzymes: α-G1P and the cofactor TPP are converted into dTDP-D-glucose by the first enzyme, glucose-1-phosphate thymidilyltransferase (RmlA; EC 2.7.7.24). Then, dTDP-Dglucose is metabolized into dTPD-4-keto-6-deoxy-D-glucose catalyzed by the enzyme dTDP-D-glucose-4,6-dehydratase (RmlB; EC 4.2.1.46). This product is converted into dTDP-4-keto-L-rhamnose through epimerization reaction catalyzed by the enzyme dTDP-4-dehydrorhamnose-3,5-epimerase (RmlC; EC 5.1.3.13). In the last step, dTDP4-keto-L-rhamnose is reduced to dTDP-L-rhamnose via dTDP-4-dehydrorhamnosereductase (RmlD; EC 1.1.1.133) (Fig. 4.4). The identification and biochemical characterization of genes involved in the formation of nucleotide-sugar precursors in other sphingans like gellan gum, S-88 and diutan gum have been performed and the corresponding genes exhibit high similarities. Based on the similarities, genes that take part in the synthesis of nucleotide-sugar precursors in welan gum formation have been annotated [23, 24]. The pgmG gene (1386 bp) has been identified in contig21 (ALBQ01000021.1) and exhibits a nucleotide identity of about 90.98 % to the known pgmG involved in the sphingan biosynthesis. As in gellan biosynthetic pathway, PgmG protein might have both PGM and phosphomannomutase (PMM) activities [25]. This bifunctional protein might play a vital role in welan gum biosynthesis because it represents a branch point in carbohydrate metabolism: α-G6P enters catabolic processes to produce energy and reducing power, while α-G1P is a precursor of nucleotide-sugar in the synthesis of many polysaccharides. The ugpG gene (870 bp) and the ugdG (1314 bp) gene are in contig11 (ALBQ01000011.1) and show high similarity to the known ugpG and ugdG

| 85

4.4 Biosynthetic pathway of welan gum

Glucose PQQ

Gluconate PQQH2

Glucose ATP

Gluconate ATP

ADP UTP

PgmG

α-G1P

α-G6P

RmlA

UgpG UDP-Glc

CO2 PMI

α-M6P

dTDP-Glc

β-F6P

R5P

NADPH

UMP

RmlC

IDP-Glc

dTDP-Rha

H2O

KDPG

α-M1P

NADP+

GTP

RmlD UDP-GlcA

6-P-G NADPH,H+

E-4-P

RmlB IP NAD(P)H,H+ UgdG

NAD(P)H,H+

GAP Pi

GDP-Man

2NADH,H+ 2ATP,CO2 AcCoA UDP

IDP

dTDP

Welan

GDP

ATP

Fig. 4.4: Biosynthetic pathway of sugar-nucleotides for welan gum synthesis in Alcaligenes sp. CGMCC2428 (modified according to Fig. 3 in reference [13] with kind permission of Springer Science and Business Media). Abbreviations: α-G6P: α-glucose-6-phosphate; β-F6P: βfructose-6-phosphate; R-5-P: ribose-5-phosphate; GAP: glyceraldehyde-3-phosphate; 6-P-G: 6-phosphogluconate; KDPG: 2-keto-3-deoxy-6- phosphogluconate; Ac-CoA: acetyl coenzyme A; E-4-P:erythrose-4-phosphate; α-G1P: α-glucose-1-phosphate; α-M6P: α-mannose-6-phosphate; αM1P: α-mannose-1-phosphate; UDP-Glc: UDP-glucose; UDP-GlcA: UDP-glucuronic acid; dTDP-Glc: dTDP-glucose; dTDP-Rha: dTDP-rhamnose; GDP-Man: GDP-mannose; IP: isoprenoid phosphate; IDP: isoprenoid pyrophosphate.

(about 88.97 % and 91.02 %), respectively. Like gel, sps, and dps clusters, the four-gene rml cluster including rmlABCD are arranged on the wel cluster of Sphingomonas sp. ATCC31555. All these genes also exhibit high identities to the known corresponding genes in other annotated sphingan clusters (in nucleotide sequences of 83–94 % and in amino acids of 88–97 % similarity). Thus, these genes are highly conserved in all annotated sphingan clusters.

4.4.2 Assembly of the tetrasaccharide repeating unit In general, the tetrasaccharide repeating units of different sphingans are assembled on activated lipid carriers catalyzed by specific glycosyltransferases through sequential transfer of sugars from the activated sugar precursors to specific ac-

86 | 4 Biological production of welan gum

ceptor molecules. Four glycosyltransferases have been elucidated in the assembly of the tetrasaccharide repeating unit. Due to high sequence identity between welB, welK, well, welQ and the equivalent genes in gel, sps, and dps clusters, the four genes were proposed to encode the four related glycosyltransferases [23, 24]. The first glycosyltransferase, glucosylisoprenylphosphate transferase (encoded by welB) transfers α-G1P from UDP-glucose to the C55 -isoprenylphosphate lipid carrier and forms glucose-isoprenylpyrophosphate. The second glycosyltransferase is glucosyl-α-pyrophosphorylpolyprenol (PPL)-1,4-glucuronosyl-transferase encoded by welK, responsible for the transfer of the second sugar, glucuronic acid, from UDP-glucuronic acid into the lipid-linked glucose intermediate. The third enzyme glucosyl-glucuronosyl-transferase that catalyzes the addition of the second D-glucose sugar is proposed to be encoded by welL and the fourth enzyme responsible for the addition of last sugar, rhamnose, is putatively encoded by welQ. Then to complete the assembly of the tetrasaccharide repeat, the substituent acetate and glycerate groups will be added to the second glucose. The genes and enzymes involved in these steps still remain unknown for all sphingans except gellan. Harding et al. [26] identified an acetyltransferase by generating an acetyltransferasedeficient mutant strain of Sphingomonas elodea that produced non-acetylated gellan, but this gene was not adjacent to the gel clusters.

4.4.3 Polymerization and export The mechanistic details of polymerization, chain-length regulation and export of sphingans after assembly of the tetrasaccharide repeating unit are still poorly understood. In the current model, the polymerization of the repeat units is thought to be in the general Wzx/Wzy-dependent pathway. For welan gum, the proposed polymerization and export process is as follows: the lipid-linked repeating units are translocated from the cytoplasmic side to the periplasmic side of the inner membrane by a Wzx-like translocase encoded by welS. Then the polymerization of the repeat units is completed by a Wzy-like-polymerase encoded by welG through transfer of the nascent polymer from its lipid carrier to the reducing end of the new lipid-linked repeat unit. The chain length of the synthesized welan gum is regulated by enzymes of the family of protein polysaccharide co-polymerases (PCPs) encoded by welC and welE in the periplasmatic space. Export of welan gum is predicted to be realized by integral outer membrane protein (encoded by welD) belonging to the outer membrane auxiliary (OMA) family [24]. The predicted protein encoded by welJ with high similarity to GelJ was proposed to be involved in the secretion and transport process because of its similarity to the AAA-superfamily of ATPases (CD0009) and ABC transporter proteins (Pfam00005). The atrB and atrD proteins show similarity to ABC-type transporter and membrane fusion proteins of type I secretion system, respectively [1]. It is possible that genes welM and welN are involved in protein folding and export based on their similarity to

4.4 Biosynthetic pathway of welan gum |

87

protein EpsH and EpsI that are proposed to form a protein export sorting system in methanolan biosynthesis [1, 27]. A clear function of WelI is not specified according to its homologous protein GelI. A deletion of gelI leads to lower gellan gum titers without composition change. The sequence analysis indicates its N-terminal lipoprotein attachment site and weak homology to peptidylprolyl cis–trans isomerases involving in protein folding. These results suggest that gelI might play a role in gellan secretion [1, 27].

4.4.4 Regulation of welan gum biosynthesis In gellan gum biosynthesis, the protein encoded by gelA is homologous to sensor kinase and response regulatory proteins. It might regulate the gellan biosynthesis [1]. welA, a homologous version of gelA (identity on protein level = 85 % and nucleotide level = 78 %), is identified by screening the draft genome of Sphingomonas sp. ATCC 31555 and proposed to encode the regulator for welan gum biosynthesis.

4.4.5 Enzymes in other process By comparison with the gel, sps and dps clusters, besides the above genes, there are other genes that may be involved in the biosynthesis of welan gum but with unproved function. The uncharacterized open reading frames urf such as urf31, urf31.4 and urf34 are proposed to be involved in the transfer of side chains or in the assembly process in other clusters especially in the dps cluster [28]. The genes urf31, urf31.4 and urf34 were also found in the wel cluster. Urf31 in the dps cluster presents some features of glycosyltransferase group 2 enzymes, such as the DXD consensus sequence (amino acid 111–113) and the aspartate residue essential for activity in other β-glycosyltransferases. This protein is predicted to be involved in addition of the rhamnosyl side chain by the deletion of urf31 in Sphingomonas ATCC 53159 which resulted in diutan with less or even without rhamnosyl side chains. The function of urf31.4 and urf34 are still unknown to date. According to the corresponding genes in other sphingans cluster, the possible function of gene welF remains unknown. Failed mutation of its homologous protein GelF indicates that either this mutation was lethal or had a polarity effect on downstream genes [1, 27]. Although the wel cluster responsible for welan biosynthesis has been predicted, the function of each gene still needs to be proved in further experiments and the bottlenecks of the biosynthetic pathway are still unclear.

88 | 4 Biological production of welan gum

4.5 Engineering approaches for improvement of sphingan production As gene clusters for sphingans synthesis have been revealed, metabolic engineering provides a promising strategy to improve their production or obtain modified sphingans. Several patents, mostly kept by Kelco Company or Shin-Etsu Bio, Inc., have been involved in genetic engineering of the different sphingan-producing strains. Several strategies in engineering Sphingomonas have been applied. One is pathway engineering to optimize the metabolic flux to sphingan synthesis. In gellan gum production, it has been found that poly-β-hydroxybutyric acid (PHB) is an energy storage product and a gellan-competing product at the optimal fermentation conditions for gellan gum synthesis. Baird and Cleary [29] tried to use PHB-deficient strains obtained by random mutagenesis to block PHB synthesis and enhance gellan gum production, but no positive effect on gellan gum synthesis was observed. Vartak et al. [30] attempted to increase its production by enhancing the low sugar conversion rate (40–50 %) by interrupting zwf gene encoding glucose-6-P-dehydrogenase in Sphingomonas elodea ATCC 31461 to divert carbon flow toward gellan gum synthesis. Similarly, the results showed that the production of gellan gum did not significantly increase. Another strategy is the overexpression of genes in the corresponding clusters that are involved in sphingan synthesis. Although the individual overexpression of the gene pgmG and ugpG led to an increase of enzyme-specific activities in the strain Sphingomonas elodea ATCC 31461, no significant enhancement of gellan gum production was observed [31]. However, when both pgmG and ugdG were overexpressed in Sphingomonas elodea ATCC 31461, its gellan gum production increased by 20 % and the biopolymer showed higher viscosity [25]. A similar result was observed by multicopy expression of pgm and the multiple sps genes required for assembly of the carbohydrate repeat unit in Sphingomonas sp. ATCC 21423 for the sphingan S-7 production. A sixfold increase of Pgm activity resulted in a small percentage increase in glucose conversion to the S-7 but multiple sps genes caused a 20 % increase in the S-7 yield and a larger increase in culture viscosity that was associated with a higher ratio of rhamnose to glucose residues in the secreted polymer compared with that in the original S-7 polymer [32]. However, in contrast to the negligible effect of overexpression of pgm on gellan and S-7 synthesis, overexpression of the pgm gene in the newly identified strain Sphingomonas sanxanigenens resulted in a 17 ± 0.3 % increase in sphingan production compared with the wild type [33]. These results might indicate different mechanisms for regulation of nucleotide-sugar precursor synthesis within the Sphingomonas family. Coleman et al. [28] tested the ability of four plasmids containing different genes from the dps cluster to enhance the production of diutan gum. They found that the four recombinant strains harboring the four plasmids produced a little more diutan gum than wild-type strain, Sphingomonas ATCC 53159 (S657). However, the recombi-

4.6 Applications of welan gum |

89

nant strain with the plasmid containing 20 of the 24 biosynthetic genes including the genes for four glycosyltransferases in the assembly of the tetrasaccharide repeating unit, the four-gene cluster for dTDP-rhamnose synthesis, genes for polysaccharide secretion and the putative genes for polymerase and lyase, led to a significant increase in EPS viscosity relative to its increase in yield. Besides introduction of various DNA segments from the sphingan cluster to enhance the production and viscosity of sphingans, deletion of genes in their biosynthetic pathway might also be used to obtain sphingans with special properties. Mutants of diutan- and gellan-producing strains in which dpsMNR/gelMNR and dpsL/gelL were deleted favored slime-forming polysaccharide production, resulting in higher centrifugal yields of biomass compared with the original strains. This method led to facilitated downstream processes together with enhanced EPS production, and optimized mass and oxygen transfer rates during the fermentation process [34]. Although these results are promising, the success of genetic engineering approaches still requires more detailed understanding of sphingan biosynthesis. Besides the genetic engineering approaches, Schmid et al. [24] think that protein engineering also shows great potential to enhance the property of EPS by modifying their structures. For example, protein engineering of glycosyltransferases will broaden the spectra of sugar donors and acceptors and might expand the portfolio of polysaccharide variants. Successful engineering of glycosyltransferases might result in EPS with varied monomer compositions and novel properties. Engineering of polymerases might realize the control of MW of EPS and the design of modified repeat units with elongated or truncated side chains or altered substituents. These different EPS will be beneficial for several fields of applications. As well as the elucidation of the wel cluster, genetic engineering and protein engineering will be efficient tools for welan gum biosynthesis.

4.6 Applications of welan gum Due to the good thermal stability and excellent rheological properties of welan gum, it has great potential to be widely applied in cement, printing ink, petroleum, food and other industries. In this section, we will discuss its possible applications.

4.6.1 Cement systems Welan gum can be used as a good cement additive because even in low concentration it can reduce fluid loss of the cement compositions, increase the suspension properties of cement suspension. Allen et al. [35] find that when comprised 0.01 to 0.9 % welan gum by weight of dry cement, the cement composition exhibits many improved features including workability, air entrainment, suspension of aggregates, sag resis-

90 | 4 Biological production of welan gum

tance, flow characteristics and resistance to water loss. What is more important is that these improvements can be retained at elevated temperatures higher than 93 °C. These properties broaden the application of cements. Sakata et al. [36] have made a liquefied viscosity welan gum agent with polycarboxylic type of AE superplasticizer. When welan gum is added to AE superplasticizer and agitated well, the particles of welan gum swell in the superplasticizer, leading to a stable suspension without much viscosity increase. Welan gum can prevent shaft lining collapse and make it easy to cut gravel when it is used as a fluid loss agent in oil-well cement. Plank et al. [37] investigated the fluid loss workability of a CaAMPS® -N,N-dimethylacrylamide copolymer (CaAMPS® co-NNDMA) supplemented with welan gum at 80 °C. The effectiveness of CaAMPS® co-NNDMA is reduced by welan gum by competing with CaAMPSV® -co-NNDMA for adsorption sites on the cement surface. It prevents the adsorption of CaAMPSV® -coNNDMA or through displacement of already adsorbed CaAMPSV® -co-NNDMA fluid loss additive molecules from the surface of cement. This is attributed to the carboxylate functionalities of welan gum which are much stronger anchor groups than the sulfonate groups of CaAMPSV® -co-NNDMA. Furthermore, due to its adsorption effect, welan gum has a retarding effect on tricalcium aluminate-gypsum hydration, a partial system of ordinary Portland cement (OPC) hydration [38].

4.6.2 Oil industry In enhanced oil recovery (EOR) processes, water-soluble polymers have been used for polymer flooding. As a high molecular extracellular polysaccharide, welan gum is expected to be used in the oil industry as a novel oil recovery agent because of its excellent rheological properties. Even at low concentration (5 %), welan gum solutions increase the viscosity of water-based drilling fluids for the displacement of oil from the reservoir [39]. Crosslinked welan gum (with phenolic resins or mixtures of phenol and aldehyde compounds) can also form a gel in low salinity brines, control permeability of subterranean strata and increase the viscosity of drilling fluids. Recently, Xu et al. [6] find that compared with the commonly-used water-soluble polymer xanthan gum, the oil displacement efficiency of welan gum is much higher in the simulated flooding experiment. In a further study, welan gum aqueous solutions present a higher viscoelasticity than that of xanthan gum in the same conditions though its MW is lower. The possible reason is that adjacent double helices of welan gum form a stable network structure in a zipper model while that of xanthan gum is just a transient one. Gao [40] observes that welan gum keeps high viscosity under high concentration of sodium ion and calcium ion, high temperature and long-term heating. Core flooding tests show that welan gum does not significantly reduce residual oil saturation, but reduces the time to reach residual oil saturation. Both results show that welan gum has great potential to be applied in EOR.

4.7 Future perspectives |

91

Besides, welan gum can also be applied in many other oil field operations such as hydraulic fracturing, well completion, well workover, wellbore clean up. It has been used as a spacer fluid in oil-well drilling. Because welan gum fluids are pumped down the hole through straight and coiled tubing, Asubiaro and Shah [41] investigate the hydraulic characteristics of welan gum fluids at different concentrations in straight and coiled tubing. They especially put emphasis on the influence of welan gum concentration and coiled tubing curvature ratio and find that increasing coiled tubing curvature ratio brings about higher friction loss for all welan gum fluids investigated.

4.6.3 Other potential applications Due to its good thickening property that is not affected by temperature and pH, welan gum has great potential to be applied in the food industry. It can be used as an ingredient in bakery products, dairy products like ice creams and yogurt, jellies and other high sugar systems, beverages like citric acid based drinks, salad dressings and various desserts [5]. For example, fat is used as one main ingredient for biscuits and other foods to give the product a good structural layer, adjust the flavor of the product and to ensure that the product has a good degree of porosity. However, it also brings high calories to the product. The application of welan gum in foods that need further processing (like frying) can produce a low calorie product by reducing its adsorption on fat. Besides, it has been reported that the polysaccharide can enhance food flavor and satisfy the needs of the consumer. In the production of acidic dairy products, welan gum as the protective colloid can eliminate protein flocculation of milk products and improve their tastes. It can also be used as a stable thickener in ice cream production at low amounts. Japan has allowed the application of welan gum as a food additive in the food industry since 1996. However, there is no experimental data on human subject research to prove the safety of welan gum until now. Welan gum also plays an important role in the ink industry because of its Nonnewtonian pseudoplastic fluid characteristics. The addition of welan gum provides shear-thinning flow characteristics to the ink. Ink with welan gum is a viscous liquid but becomes a thin, readily flowable liquid at shear rates generated in the writing process. Furthermore, welan gum keeps the ink stable at high temperatures (greater than 100 °C) in the processing, storage, and transport process or by the user [42].

4.7 Future perspectives Over the last years, important advances have been made in the production of welan gum. Prediction of the wel cluster responsible for welan biosynthesis deepens the understanding of the possible biosynthetic pathway of welan gum. However, the function of each gene needs to be proved in further experiments and the bottlenecks of the

92 | 4 Biological production of welan gum

pathway are still unclear. Although the production of welan gum has improved by optimization of the fermentation conditions and strain breeding, the conversion rate of glucose and the yield of welan gum are much lower than that of xanthan gum while its production cost is much higher. Further efforts are needed to enhance its yield and the glucose conversion rate, and to reduce the production cost. One possible way is utilization of effective low cost substrates such as agricultural waste or dairy waste in the fermentation process. Another way is metabolic engineering of the biosynthetic pathway of welan gum based on the wel cluster. Many attempts have been made to broaden the application of welan gum. Welan gum has great potential to be applied in self-consolidating concrete, the oil, food and ink industries. Research is required to expand its use in other areas like the pharmaceutical industry and pesticides. In the pharmaceutical industry, it might increase corneal contact time with the drug and could be used in sustained-release control of tablets.

References [1]

Fialho AM, Moreira LM, Granja AT, Popescu AO, Hoffmann K, Sá-Correia I. Occurrence, production, and applications of gellan: current state and perspectives, Appl Microbiol Biotechnol, 2008, 79, 889–900. [2] Jay AJ, Colquhoun IJ, Ridout MJ, Brownsey GJ, Morris VJ, Fialho AM, Leitão JH, Sá-Correia I. Analysis of structure and function of gellans with different substitution patterns, Carbohydr Polym, 1998, 35, 179–88. [3] Jansson PE, Widmalm G. Welan gum (S-130) contains repeating units with randomly distributed L-mannosyl and L-rhamnosyl terminal groups, as determined by FABMS, Carbohydr Res, 1994, 256, 327–30. [4] Chandrasekaran R, Radha A, Lee EJ. Structural roles of calcium ions and side chains in welan: An X-ray study, Carbohydr Res, 1994, 252, 183–207. [5] Kang KS, Veeder GT. Heteropolysaccharide S-130. Merck & Co., Inc., US patent, 4342866, 1981. [6] Xu L, Xu G, Liu T, Chen Y, Gong H. The comparison of rheological properties of aqueous welan gum and xanthan gum solutions, Carbohydr Polym, 2013, 92,516–22. [7] Xu H, Li S, Guo CJ, Ouyang PK. One strain of Alcaligenes sp. and its application in welan gum preparation. CN patent, 200610088356.0, 2006. [8] Li S, Xu H, Li H, Guo C. Optimizing the production of welan gum by Alcaligenes facalis NX-3 using statistical experiment design, Afr J Biotechno, 2010, 9, 1024–1030. [9] Ma T, Li GQ, Li DP, Liang FL, Liu RL. One strain of Sphingomonas sp. TP-5 and its application in welan gum production. CN patent, 200910069960.2, 2009. [10] Li H, Xu H, Li S, Xu H, Guo CJ, Ying HJ, Ouyang PK. Strain improvement and metabolic flux modeling of wild-type and mutant Alcaligenes sp. NX-3 for synthesis of exopolysaccharide welan gum, Biotechnol Bioproc E, 2010, 15, 777–84. [11] Zhu P, Chen X, Li S, Xu H, Dong S, Xu Z, Zhang Y. Screening and characterization of Sphingomonas sp. mutant for welan gum biosynthesis at an elevated temperature, Bioprocess Biosyst Eng, 2014, 37, 1849–58. [12] Zhao Y, Chen F, Li JK, Liao Bin, Tu YG. Fermentation processing of microbial polysaccharide welan gum, Food Science, 2010, 31, 219–23 (in Chinese).

References

|

93

[13] Li H, Xu H, Xu H, Li S, Ouyang PK. Biosynthetic pathway of sugar nucleotides essential for welan gum production in Alcaligenes sp. CGMCC2428, Appl Microbiol Biotechnol, 2010, 86, 295–303. [14] Li H, Xu H, Li S, Feng XH, Ouyang PK. Optimization of exopolysaccharide welan gum production by Alcaligenes sp. CGMCC2428 with Tween-40 using response surface methodology, Carbohydr Polym, 2012, 87, 1363–8. [15] Li H, Li S, Xu H, Chen XY, Ouyang PK. Improvement of welan gum production and redistribution of metabolic flux under pH control process in Alcaligenes sp. CGMCC2428, Biotechnol Bioproc E, 2013, 18, 399–405. [16] Banik RM, Santhiagu A. Improvement in production and quality of gellan gum by Sphingomonas paucimobilis under high dissolved oxygen tension levels, Biotechnol Lett, 2006, 28, 1347–50. [17] Giavasis I, Harvey LM, McNeil B. The effect of agitation and aeration on the synthesis and molecular weight of gellan in batch cultures of Sphingomonas paucimobilis, Enzyme Microb Technol, 2006, 38, 101–8. [18] Li H, Xu H, Xu H, Li S, Ying HJ, Ouyang PK. Enhanced welan gum production using a two-stage agitation speed control strategy in Alcaligenes sp. CGMCC2428, Bioprocess Biosyst Eng, 2011, 34, 95–102. [19] Li H, Xu H, Li S, Feng XH, Xu H, Ouyang PK. Effects of dissolved oxygen and shear stress on the synthesis and molecular weight of welan gum produced from Alcaligenes sp. CGMCC2428, Process Biochem, 2011, 46, 1172–8. [20] O’Neill MA, Selvendran RR, Morris VJ, Eagles J. Structure of the extracellular polysaccharide produced by the bacterium Alcaligenes (ATCC 31555) species, Carbohydr Res, 1986, 147, 295–313. [21] Cannon JJ. Polysaccharide isolation process. US Patent, 923924, 1988. [22] Kaur V, Bera MB, Panesar PS, Kumar H, Kennedy JF. Welan gum: microbial production, characterization, and applications, Int J Biol Macromol, 2014, 65, 454–61. [23] Wang X, Tao F, Gai Z, Tang H, Xu P. Genome sequence of the welan gum-producing strain Sphingomonas sp. ATCC 31555, J Bacteriol, 2012, 194, 5989–90. [24] Schmid J, Sperl N, Sieber V. A comparison of genes involved in sphingan biosynthesis brought up to date, Appl Microbiol Biotechnol, 2014, 98, 7719–33. [25] Sá-Correia I, Fialho AM, Videira P, Moreira LM, Marques AR, Albano H. Gellan gum biosynthesis in Sphingomonas paucimobilis ATCC 31461: genes, enzymes and exopolysaccharides production engineering, J Ind Microbiol Biotechnol, 2002, 29, 170–6. [26] Harding N, McQuown J, Patel YN. Mutant strain of Sphingomonas elodea which produces nonacetylated gellan gum. CP Kelco U.S., Inc, Delaware. US patent, 0100078 A1, 2003. [27] Harding NE, Patel YN, Coleman RJ. Organization of genes required for gellan polysaccharide biosynthesis in Sphingomonas elodea ATCC 31461, J Ind Microbiol Biotechnol, 2004, 31, 70–82. [28] Coleman RJ, Patel YN, Harding NE. Identification and organization of genes for diutan polysaccharide synthesis from Sphingomonas sp. ATCC 53159, J Ind Microbiol Biotechnol, 2008, 35, 263–74. [29] Baird JK and Cleary JM. PHB- Free Gellan Gum Broth. Merck & Co., Inc., US Patent, 5300429, 1994. [30] Vartak NB, Lin CC, Cleary JM, Fagan MJ, Saier Jr MH. Glucose metabolism in Sphingomonas elodea: pathway engineering via construction of a glucose-6-phosphate dehydrogenase insertion mutant, Microbiology, 1995, 141, 2339–50. [31] Videira PA, Cortes LL, Fialho AM, Sa-Correia I. Identification of the pgmG gene, encoding a bifunctional protein with phosphoglucomutase and phosphomannomutase activities, in the

94 | 4 Biological production of welan gum

[32]

[33]

[34] [35] [36] [37]

[38] [39] [40] [41] [42]

gellan gum producing strain Sphingomonas paucimobilis ATCC 31461, Appl Environ Microbiol, 2000, 66, 2252–8. Thorne L, Mikolajczak MJ, Armentrout RW, Pollock TJ. Increasing the yield and viscosity of exopolysaccharides secreted by Sphingomonas by augmentation of chromosomal genes with multiple copies of cloned biosynthetic genes, J Ind Microbiol Biotech, 2000, 25, 49–57. Huang H, Li X, Wu M, Wang S, Li G, Ma T. Cloning, expression and characterization of a phosphoglucomutase/phosphomannomutase from sphingan-producing Sphingomonas sanxanigenens, Biotechnol Lett, 2013, 35, 1265–70. Harding NE, Yamini P, Coleman RJ. Targeted gene deletions for polysaccharide slime formers. Cp Kelco U.S., Inc, US patent, 8759071 B2, 2006. Allen F L, Best G H, Lindroth TA. Welan gum in cement compositions. Merck & Co., Inc. US patent, 4963668, 1990. Sakata N, Yanai S, Yokozeki K, Maruyama K. Study on new viscosity agent for combination use type of self-compacting concrete, J Adv Concr Technol, 2003, 1, 37–41. Plank J, Lummer NR, Dugonjic-Bilic F. Competitive adsorption between an AMPSV® -based fluid loss polymer and Welan gum biopolymer in oil well cement, J Appl Polym Sci, 2010, 116, 2913–9. Ma L, Zhao Q, Yao C, Zhou M. Impact of welan gum on tricalcium aluminate-gypsum hydration, Mater Char, 2012, 64, 88–95. Takahashi K, Sugiura T, Takeuchi T. Oil absorption retarder. Kibun Food Chemifa Co., Ltd, US patent 6497910, 2002. Gao CH. Potential of Welan gum to enhance oil recovery, J Petrol Explor Prod Technol, 2015, 5, 197–200. Asubiaro A, Shah NS. Rheological and hydraulic properties of welan gum fluids in straight and coiled tubings, J Fluids Eng, 2008, 130, 081506. Wang A, Chadwick BW. Ink composition. BIC Corporation, US patent, 5769931, 1998.

Han Wei, Tang Junhong, and Li Yongfeng

5 Utilization of food waste for fermentative hydrogen production 5.1 Introduction Current imperative global issues such as petroleum depletion and global warming are leading to new developments in fuel markets all over the world [1]. Interest in the development of renewable energy to reduce the reliance on fossil fuels and achieve sustainable development in energy consumption is increasing [2, 3]. Hydrogen is a promising alternative to fossil fuels because it is clean and renewable [4]. The energy yield of hydrogen is 122 kJ/g, which is 2.75 times higher than fossil fuel [5]. Moreover, hydrogen can be directly used to produce electricity via fuel cells [6]. Therefore, hydrogen is considered to be a promising energy carrier of the future. The best-known industrial ways of hydrogen production are steam reformation of natural gas, coal gasification and splitting water with electricity [7, 8]. However, these industrial processes could also release carbon dioxide and other greenhouse gases and pollutants as byproducts [9]. Recently, biological hydrogen production has attracted considerable attention since it could deal with the conversion of low cost residues or organic waste/wastewater to hydrogen [10, 11]. Biological hydrogen production processes are considered to be more environmentally friendly and less energy intensive compared to thermochemical and electrochemical processes [12]. Generally, biological hydrogen production can be divided into two categories: photosynthesis and dark fermentation [13]. Dark fermentation seems to be a more feasible biotechnology for hydrogen production than photosynthesis due to lower energy consumption and no light limitation [14]. However, the low hydrogen production rate and high cost are the dominant obstacles for large-scale dark fermentative hydrogen production [15]. Utilization of raw waste/wastewater as substrate for fermentative hydrogen production (such as food waste) could effectively enhance the economic benefit which is regarded as a promising solution [16]. Food waste is a promising raw material for biofuel production because of its high organic content and availability. It mainly consists of starch, protein and fat which are good carbon sources for fermentative hydrogen production [17]. Fermentative bacteria hydrolyze and ferment carbohydrates, protein and lipids to volatile fatty acids which are then further converted into acetate, carbon dioxide and hydrogen by acetogenic bacteria [18]. Hydrogen and ATP are produced by fermentative bacteria such as Clostridium sp. during the degradation process. The limiting factor for biohydrogen production from food waste is the hydrolysis rate [19]. Kim et al. [20] found that heatpretreated food waste could accelerate the hydrolysis rate of food waste and produce high biohydrogen yield when compared to untreated food waste. Similarly, sonication

96 | 5 Utilization of food waste for fermentative hydrogen production

of food waste with heat and without inoculum was applied by Elbeshbishy et al. [21] for biohydrogen production. This research showed that pretreatment of food waste could enhance biohydrogen production efficiency and therefore can be regarded as an important parameter influencing biohydrogen production. Enzymatic hydrolysis could release nutrients (such as glucose and free amino nitrogen) from food waste with the advantage of a high hydrolysis rate and mild reaction conditions [22, 23]. Therefore, this chapter presents an updated review on dark fermentative hydrogen production from food waste. The analysis performed in the present chapter was focused on the following issues: (1) metabolic pathway of fermentative hydrogen production, (2) characteristics of food waste affecting the performance of fermentative hydrogen production, (3) pretreatment of food waste for fermentative hydrogen production.

5.2 Metabolic pathway of fermentative hydrogen production 5.2.1 Process yield and conversion efficiency The concept of conversion efficiency derives from the existence of a fermentation barrier to hydrogen production from organic substrates. If the complete conversion reaction to hydrogen is taken into account (Eq. (5.1)), it is concluded that theoretically 12 mol hydrogen could be generated from 1 mol glucose [24, 25]. C6 H12 O6 + 6H2 O → 12H2 + 6CO2

(5.1)

However, this reaction is energetically unfavorable with respect to biomass growth and would occur only with extremely low hydrogen concentration. The optimal conversion of glucose into hydrogen is limited by acetate production. As a result, one third of the theoretical hydrogen production can be achieved in practice because part of the reducing equivalents in the original substrate remains as acetate (Eq. (5.2)) [26]. C6 H12 O6 + 2H2 O → 4H2 + 2CO2 + 2CH3 COOH

(5.2)

In practice, organic intermediates act as electron scavengers, which give rise to the production of more reduced fermentation products compared to acetate, including propionate, butyrate and ethanol, with an associated decrease in the hydrogen yield. In case the butyrate fermentation pathway is established, the conversion efficiency is reduced to 2 mol H2 /mol glucose (Eq. (5.3)) [27]. C6 H12 O6 → 2H2 + 2CO2 + CH3 CH2 CH2 COOH

(5.3)

5.2 Metabolic pathway of fermentative hydrogen production

|

97

5.2.2 Metabolic pathway for fermentative hydrogen production The carbohydrate must undergo liquefaction by extracellular enzymes before being taken up by acidogenic bacteria. The rate of hydrolysis is a function of several factors, such as pH and temperature [28, 29]. After that, soluble organic components, including the products of hydrolysis, are converted into organic acids, ethanol, hydrogen and carbon dioxide by acidogens (Fig. 5.1). The products of acidogenesis are then converted into acetate, hydrogen and carbon dioxide [30].

C6H12O6

Carbohydrate 2CO2

2CO2

C6H12O6 2NAD+

2CH3CH2OH

2ADP +

4NAD 2CH3CHOOH

4NADH+H

+

2ATP

2ADP

2NADH+H+

2FdH2

2Fd

2CH3(CH2)2COOH

2NAD+ 2CH3CHOHCOOH

4NADH+H+

ADP 2ADP

2NAD+

2NADH+H+

2CH3COCOOH

2CH3COSCo 2ATP

2NADH+H+

ATP

4NAD+

2ATP

2H2

2CH3CH2COOH

Fig. 5.1: Metabolic pathway and byproducts in fermentative hydrogen production [58].

Fermentative hydrogen production is carried out by anaerobic acidogenic bacteria with highly diverse fermentation characteristics and hydrogen production capabilities. Performance of fermentative hydrogen production depends on a number of parameters, such as pH, temperature, and organic loading rate [31–33]. The variations of parameters would lead to various microbial communities which finally result in diverse fermentation types [34]. There are four main fermentation types in the anaerobic acidogenesis, namely acetate type fermentation, butyrate type fermentation, ethanol type fermentation and propionate type fermentation [35–37]. Many microbial communities exhibit acetate fermentation with acetate as the major product (Eq. (5.4)). The major products of propionate type fermentation are propionate and acetate (Eq. (5.5)), while the products of ethanol type fermentation are ethanol and acetate (Eq. (5.6)).

98 | 5 Utilization of food waste for fermentative hydrogen production

As for butyrate type fermentation, butyrate and acetate are the primary fermentation products (Eq. (5.7)). C6 H12 O6 + 2H2 O → 2CH3 COOH + 4H2 + 2CO2

(5.4)

C6 H12 O6 → 2CH3 CH2 COOH + 2CO2

(5.5)

C6 H12 O6 + H2 O → CH3 CH2 OH + CH3 COOH + 2H2 + 2CO2

(5.6)

C6 H12 O6 → CH3 (CH2 )2 COOH + 2H2 + 2CO2

(5.7)

Equations (5.4)–(5.7) show that hydrogen is produced from acetate, butyrate and ethanol type fermentations. Propionate type fermentation could not generate hydrogen. However, propionate type fermentation is concurrent with other fermentation types capable of producing hydrogen in a mixed microbial community [38, 39]. Therefore, hydrogen could also be generated from anaerobic fermentation when the production of propionate is still high. In many papers, butyrate type fermentation is considered as the most common pathway for fermentative hydrogen production. Relative research about ethanol type fermentation remains deficient. Based on the equilibrium of the NADH/NAD+ ratio inside the bacteria cell, Ren et al. [40] proposed that the ethanol type pathway induced at pH 4.5 is a better and more stable metabolic pathway than the butyrate type pathway induced at pH 5.0. Although the theoretical yield of hydrogen is 2 mol hydrogen/mol glucose in butyrate type fermentation (Reaction 7), which is same as that of the ethanol type fermentation (Reaction 6), butyrate type fermentation lacks the stability for NADH accumulation because part of the produced NADH can be utilized rapidly by cellular synthesis or converted to hydrogen and NAD+ under the presence of acetyl-CoA [41]. So, the butyrate production pathway has the potential to change to the butanol production pathway, where hydrogen may be consumed [42, 43]. Conversely, it can be inferred from Fig. 5.2 that ethanol type fermentation is able to preserve a balance of NADH + H+ /NAD+ . The carbohydrate is first degraded to glucose which is further converted to pyruvate (CH3 COCOH). Pyruvate is oxidized to CH3 COSCoA by depletion of NAD+ with molecular hydrogen and carbon dioxide generation. In order to keep sequential production of hydrogen, the metabolism product NADH + H+ must be utilized to regenerate NAD+ to compensate for equilibrium between the NADH + H+ and NAD+ by the reaction of the ethanol-acetate pathway [44]. This fermentation pathway can reduce acidic terminal products by producing neutral matter of ethanol and make the acidogenic fermentation process favorable for hydrogen production [45]. This shows that ethanol type fermentation can obtain better stability and no pH regulation was required for this fermentation during the whole operation process. Therefore, ethanol type fermentation is the optimal choice for maximum hydrogen production by mixed culture [46–48]. Despite the potential advantages, further deep studies are wort doing to examine the metabolic pathway using genetic modification of hydrogen-producing bacteria and further clarify in more detail the control strategy of ethanol type fermentation.

5.3 Biohydrogen production from food waste |

ADP C6H12O6 NAD+

HSCoA CO2

ATP

NADH +H+

HSCoA CH3COSCoA

CH3COCOH Fd

FdH2

99

2ADP 2ATP

CH3CHO NADH NAD+ +H+

CH3CH2OH NADH NAD+ +H+

CH3COOH H2 Fig. 5.2: Amended ethanol type fermentation route by acidogenic bacteria [22, 33].

5.3 Biohydrogen production from food waste There are two main ways of biological hydrogen production from carbohydrate: dark fermentation and photosynthesis [49]. The major substrates used for dark fermentation are simple sugars, such as glucose and sucrose. However, the substrates used for photosynthesis are organic acids, such as acetate and butyrate [50]. Dark fermentation is considered to be a more feasible biotechnology for hydrogen production than photosynthesis due to lower energy consumption and no light limitation [51]. However, the low hydrogen production rate and high cost are the dominant obstacles for large-scale dark fermentative hydrogen production [52]. Utilization of raw waste/wastewater as substrate for fermentative hydrogen production (such as food waste) could effectively enhance the economic benefit which is regarded as a promising solution [53]. Food waste is one of the most severe environmental problems all over the world [54]. Over a billion tons of food waste is generated per year which accounts for 33 % of annual global food production [55]. Therefore, disposal and utilization of food waste is becoming one of the major global challenges. Food waste consists mainly of starch and protein which make food waste an economical source for biofuel production [56]. Utilization of food waste for hydrogen production could not only solve the food waste problem, but also produce an alternative energy source simultaneously [16, 57]. However, nutrients stored in food waste are in the form of macromolecules (such as starch and protein) which have to be broken into utilizable forms (glucose and free amino nitrogen) before being utilized by microorganisms for fermentative hydrogen production [24, 58]. Generally, there are two main stages in fermentative hydrogen production from food waste (hydrolysis and fermentation). Separate hydrolysis and fermentation is the process in which food waste is first hydrolyzed by pretreatments to obtain micromolecules. Then, the nutrients solution is subjected to dark fermentation for hydrogen production. The hydrolysis stage of complex substrate is the rate limiting step in most of the bioprocesses [59]. However, hydrolysis of food waste in a separate process could overcome this problem. The operating conditions in pretreatment can be optimized to get the maximum food waste to nutrients solution conversion rate [60].

100 | 5 Utilization of food waste for fermentative hydrogen production

5.3.1 Carbohydrate Food waste is considered to be a suitable substrate for fermentative hydrogen production since it is rich in carbohydrate. The carbohydrate has to be hydrolyzed by hydrolytic bacteria to produce simple sugars, such as glucose and sucrose, before being utilized as substrate for fermentative hydrogen production. The product of carbohydrate hydrolysis mainly depends on the microorganisms present in the culture broth. The speed of carbohydrate hydrolysis is faster than that for lipid and protein. Lay et al. [61] indicated that the yield of hydrogen production from carbohydrate rich substrate is 20 times higher than using lipid and protein rich substrate. Sagnak et al. [62] applied both acid and heat treatments to get monomeric sugar for fermentative hydrogen production. Han et al. [63] added glucoamylase and protease to the food waste before hydrogen production to increase the efficiency of starch and protein hydrolysis.

5.3.2 Fats Oils are sources of lipids in food waste [16]. The presence of lipids in anaerobic fermentation could lead to flotation and mass transfer problems. The process of fermentative hydrogen production from lipid hydrolysis would be slower than carbohydrate hydrolysis because of the ability of hydrogenotrophic methanogens to consume hydrogenproducing bacteria [34]. Therefore, it is acknowledged that lipids are not suitable to be utilized as the sole substrate for fermentative hydrogen production.

5.3.3 Protein Food waste contains significant amounts of protein which are polypeptides formed by joining covalently linked amino acid [53]. The hydrolysis of protein is performed to produce amino acids by proteases excreted by microorganisms. Then, the amino acids are further utilized to generate volatile fatty acids, carbon dioxide and hydrogen. The speed of protein hydrolysis is slower than carbohydrate and lipid hydrolysis. Therefore, it is not suitable to use protein as sole substrate for fermentative hydrogen production.

5.4 Pretreatment of food waste for fermentative hydrogen production Depending on the food waste structure, pretreatment could be applied in single or multiple steps, including physical, chemical and enzymatic pretreatments. Physical pretreatment is related to size reduction or the contribution of a physical force to de-

5.4 Pretreatment of food waste for fermentative hydrogen production

|

101

compose the food waste structure. Chemical pretreatment is usually applied in severe acidic or alkaline conditions. Enzymatic pretreatment could be accomplished at ambient operation conditions with higher conversion rate and yield.

5.4.1 Physical pretreatment Physical pretreatment of food waste could reduce the size of food waste by physical forces without chemicals or microorganisms [51]. Comminution is the most common physical pretreatment. The main objective of physical pretreatment is to improve the available surface area by reducing the substrate size. It enables a more efficient chemical or microbial hydrolysis of the substrate matrix and decreases hydrolytic enzyme limitations. Physical pretreatment is one of the most common ways applied in fermentative hydrogen production from food waste. Reducing the size of food waste by mechanical comminution is an energy intensive process which could be achieved by different devices, such as shredders and grinders.

5.4.2 Chemical pretreatment Chemical pretreatment is the process to depolymerize the food waste using chemicals [28]. The goal of chemical pretreatment is to enable enzymatic access to fermentable sugars by breaking down the macromolecules into micromolecules. Acid and alkaline are the most commonly applied chemical pretreatment. Dilute acid hydrolysis includes HCl, H2 SO4 , and HNO3 . Dilute acid hydrolysis can be accomplished at 100–250 °C, 0.5–30 min with 0.5–3 % acid concentrations [55]. The main disadvantages of acid hydrolysis are the toxic byproducts, such as furfural, which would inhibit the performance of hydrogen production in the fermentation step.

5.4.3 Enzymatic pretreatment Food waste could be used as substrate for fermentative hydrogen production after physical or chemical pretreatment processes [10]. However, it has been observed that physical or chemical pretreatment could require intensive energy, chemicals and severe operation conditions leading to wastewater and toxic byproduct formation. Therefore, the selection of an environmental friendly and sustainable process is of great importance. Enzymatic pretreatment is regarded as an alternative option to physical and chemical pretreatment of food waste. Microorganisms, such as fungi (Aspergillus awamori and Aspergillus oryzae) and some bacteria (Clostridium thermocellum), can produce glucoamylase and protease which could degrade macromolecules (starch and protein) to release fermentable nu-

102 | 5 Utilization of food waste for fermentative hydrogen production

trients from food waste [31]. Compared to physical and chemical pretreatments, enzymatic pretreatment could operate under mild condition without toxic byproduct formation. Glucoamylase could degrade starch into glucose which can further be utilized as substrate for fermentative hydrogen production. Meanwhile, protease could hydrolyze protein into free amino nitrogen (FAN). Usually, food waste pretreatment starts with physical size reduction followed by diverse combinations of chemical and enzymatic processes. Tab. 5.1: Comparison of the performance of hydrogen production from food waste. Substrate

Microorganisms

Reactor type

H2 yield (ml H2 /g VSadded )

References

Food waste (grain, vegetables, meats and fish)

Sewage sludge

Continuous

205

[3]

Sonicated food waste

No inoculum

Batch

97

[62]

Food waste

Clostridium-rich composts

Batch

77

[61]

Food waste

Escherichia cloacae

Batch

52

[30]

Food waste

Sewage sludge

Continuous

165

[12]

Food waste

Anaerobic digester sludge

Packed-bed reactor

249

[24]

Food waste

Biohydrogenbacterium R3

Batch

294.47

[63]

VSadded : volatile solidadded .

5.5 Performance of biohydrogen production from food waste Table 5.1 summarizes the comparison of the performance of fermentative hydrogen production from food waste. Lee and Chung [35] conducted a cost analysis of hydrogen production from food waste using two-phase hydrogen/methane fermentation and suggested that the abundance and low cost of food waste makes it economically more feasible than the other sources for H2 production. Han et al. [63] developed a novel combination bioprocess of solid-state fermentation (SSF) and fermentative hydrogen production from food waste. Food waste was first utilized in solid-state fermentation by Aspergillus awamori and Aspergillus oryzae to produce glucoamylase and protease, respectively, which were used to hydrolyze food waste to obtain the food waste hydrolysate rich in glucose and free amino nitrogen (FAN). Then, the food waste hydrolysate was used as substrate for fermentative hydrogen production by heat pretreated sludge. The best hydrogen yield (52.4 ml H2 /g food waste or 294.47 ml

5.6 Prospects and challenges of fermentative hydrogen production from food waste

| 103

H2 /VSadded) was achieved at food waste mass ratio of 5 %. The proposed combination bioprocess could effectively accelerate the hydrolysis rate, improve raw material utilization and enhance hydrogen yield.

5.6 Prospects and challenges of fermentative hydrogen production from food waste Recently, fermentative hydrogen production from food waste has attracted great attention. According to the UN Food and Agriculture Organization, around 1.3 billion tons of food is wasted per year. Food waste, which is comprised mainly of starch, protein and fat, becomes a feasible source for fermentative hydrogen production. A survey was carried out to predict the development of the fermentative hydrogen production sector worldwide. It was found that China would get the largest fermentative hydrogen production market, following by the US, Japan, and India. Additional research is required to improve the efficiency of fermentative hydrogen production from food waste. It is hoped that the limitations to fermentative hydrogen production from food waste can be solved in the near future.

References [1] [2]

[3] [4]

[5]

[6] [7] [8]

[9]

Abbasi T, Abbasi SA. Renewable hydrogen: prospects and challenges, Renew Sust Energ Rev, 2011, 15, 3034–40. American Public Health Association (APHA), American Water Works Association (AWWA), Water Pollution Control Federation (WPCF). Standard methods for the examination of water and wastewater, 20th ed., 1998, Washington D.C. Chu CF, Li YY, Xu KQ, Kong HN. A pH- and temperature- phase two-stage process for hydrogen and methane production from food waste, Int J Hydrogen Energy, 2008, 33, 4739–46. Du CY, Lin SKC, Koutinas A, Wang RH, Dorado P, Webb C. A wheat biorefining strategy based on solid-state fermentation for fermentative production of succinic acid, Bioresour Technol, 2008, 99, 8310–5. Elbershbishy E, Hafez H, Dhar BR, Nakhla G. Single and combined effect of various pretreatment methods for biohydrogen production from food waste, Int J Hydrogen Energy, 2011, 36, 11379–87. Gioannis GD, Muntoni A, Polettini A, Pomi R. A review of dark fermentative hydrogen production from biodegradable municipal waste fractions, Waste Manage, 2013, 33, 1345–61. Han SK, Shin HS. Biohydrogen production by anaerobic fermentation of food waste, Int J Hydrogen Energy, 2004, 29, 569–77. Han W, Wang B, Zhou Y, Li YF, Ren NQ. Fermentative hydrogen production from molasses wastewater in a continuous mixed immobilized sludge reactor, Bioresour Technol, 2012, 110, 219–23. Han W, Liu DN, Shi YW, Tang JH, Li YF, Ren NQ. Biohydrogen production from food waste hydrolysate using continuous mixed immobilized sludge reactors, Bioresour Technol, 2015, 180, 54–8.

104 | 5 Utilization of food waste for fermentative hydrogen production

[10] Karagiannidis A, Perkoulidis G. A multi-criteria ranking of different technologies for the anaerobic digestion for energy recovery of the organic fraction of municipal solid wastes, Bioresour Technol, 2009, 100, 2355–60. [11] Ramírez-Morales JE, Tapia-Venegas E, Nemestothy N, Bakonyi P, Belafi-Bako K, Ruiz-Filippi G. Evaluation of two gas membrane modules for fermentative hydrogen separation, Int J Hydrogen Energy, 2013, 38, 14042–52. [12] Kim DH, Kim SH, Shin HS. Hydrogen fermentation of food waste without inoculum addition, Enzyme Microb Technol, 2009, 45, 181–7. [13] Lay J, Fan K, Hwang J, Chang J, Hsu P. Factors affecting hydrogen production from food waste by Clostridium-rich composts, J Environ Eng, 2005, 131, 595–602. [14] Lee DH, Chiu LH. Development of a biohydrogen economy in the United States, China, Japan, and India: With discussion of a chicken-and-egg debate, Int J Hydrogen Energy, 2012, 37, 15736–45. [15] Lee KS, Hsu YF, Lo YC, Lin PJ, Lin CY, Chang JS. Exploring optimal environmental factors for fermentative hydrogen production from starch using mixed anaerobic microflora, Int J Hydrogen Energy, 2008, 33, 1565–72. [16] Leung CCJ, Cheung ASY, Zhang AYZ, Lam KF, Lin CSK. Utilisation of waste waste bread for fermentative succinic acid production, Biochem Eng J, 2012, 65, 10–5. [17] Logan BE, Oh S, Kim IS, Van-Ginkel S. Biological H2 production measured in batch anaerobic respirometers, Environ Sci Technol, 2002, 36, 2530–5. [18] Pleissner D, Lam WC, Han W, Lau KY, Lin CSK. Fermentative polyhydroxybutyrate production from a novel feedstock derived from bakery waste, Biomed Research Int, 2014, Volume 2014, Article ID 819474. [19] Pleissner D, Lam WC, Sun Z, Lin CSK. Food waste as nutrient source in heterotrophic microalgae cultivation, Bioresour Technol, 2013, 137, 139–46. [20] Kim DH, Kim SH, Kim HW, Kim MS, Shin HS. Sewage sludge addition to food waste synergistically enhances hydrogen fermentation performance, Bioresour Technol, 2011, 102, 8501–6. [21] Elbeshbishy E, Hafez H, Dhar BR, Nakhla G. Single and combined effect of various pretreatment methods for biohydrogen production from food waste, Int J Hydrogen Energy, 2011, 36, 11379–87. [22] Ren NQ, Wang DY, Yang CP, Wang L, Xu JL, Li YF. Selection and isolation of hydrogen-producing fermentative bacteria with high yield and rate and its bioaugmentation process, Int J Hydrogen Energy, 2012, 35, 2877–82. [23] Sagnak R, Kargi F, Kapdan IK. Bio-hydrogen production from acid hydrolyzed waste ground wheat by dark fermentation, Int J Hydrogen Energy, 2011, 36, 12803–9. [24] Shin HS, Kim SH, Han SK, Kim HW, Oh SE. Current technical development in continuous H2 and CH4 production from organic waste, J Environ Eng Manage, 2006, 16(4), 217–24. [25] Show KY, Lee DJ, Tay JH, Lin CY, Chang JS. Biohydrogen production: current perspectives and the way forward, Int J Hydrogen Energy, 2012, 34, 15616–31. [26] Tapia-Venegaset E, Ramirez JE, Donoso-Bravo A, Jorquera L, Steryer JP, Ruiz-Filippi G. Biohydrogen production during acidogenic fermentation in a multistage stirred tank reactor, Int J Hydrogen Energy, 2013, 38, 2185–90. [27] Tawfik A, Salem A, El-Qelish M. Two stage anaerobic baffled reactor for biohydrogen production from municipal food waste, Bioresour Technol, 2011, 102, 8723–6. [28] Van-Ginkel SW, Oh SE, Logan BE. Biohydrogen gas production from food processing and domestic wastewaters, Int J Hydrogen Energy, 2005, 30, 1535–42. [29] Wang A, Sun D, Cao G, Wang H, Ren N, Wu WM, Logan BE. Integrated hydrogen production process from cellulose by combining dark fermentation, microbial fuel cells, and a microbial electrolysis cell, Bioresour Technol, 2011, 102, 4137–43.

References

|

105

[30] Xiao LP, Deng ZY, Fung KY, Ng KM. Biohydrogen production from anaerobic digestion of food waste, Int J Hydrogen Energy, 2013, 38, 13907–13. [31] Zhang HS, Bruns MA, Logan BE. Biological hydrogen production by Clostridium acetobutylicum in an unsaturated flow reactor, Water Res, 2006, 40, 728–34. [32] Zhang S, Lee YH, Kim TH, Hwang SJ, Effects of OLRs and HRTs on hydrogen production from high salinity substrate by halophilic hydrogen producing bacterium (HHPB), Bioresour Technol, 2013, 141, 227–32. [33] Ren NQ, Guo WQ, Wang XJ. Effects of different pretreatment methods on fermentation types and dominant bacteria for hydrogen production, Int J Hydrogen Energy, 2008, 33(16), 4318–24. [34] Das D, Verziroglu TN. Hydrogen production by biological processes: a survey of literature, Int J Hydrogen Energy, 2001, 26, 13–28. [35] Hussy I, Hawkes FR, Dinsdale R, Hawkes DL. Continuous fermentative hydrogen production from sucrose and sugar beet, Int J Hydrogen Energy, 2005, 30, 471–83. [36] Li C, Fang HHP. Fermentative hydrogen production from wastewater and solid wastes by mixed cultures, Crit Rev Environ Sci Technol, 2007, 37(1), 1–39. [37] Logan BE, Oh SE, Kim IS, Van Ginkel S. Biological hydrogen production measured in batch anaerobic respirometers, Environ Sci Technol, 2002, 36(11), 2530–5. [38] Lin CY, Lay CH. Effects of carbonate and phosphate concentrations on hydrogen production using anaerobic sewage sludge microflora, Int J Hydrogen Energy, 2004, 29(3), 275–81. [39] Wang J, Wan W. Factors influencing fermentative hydrogen production: A review, Int J Hydrogen Energy, 2009, 34(2), 799–811. [40] Ren NQ, Guo WQ, Wang XJ. Effects of different pretreatment methods on fermentation types and dominant bacteria for hydrogen production, Int J Hydrogen Energy, 2008, 33(16), 4318–24. [41] Hawkes FR, Hussy I, Kyazze G, Dinsdale R, Hawkes DL. Continuous dark fermentative hydrogen production by mesophilic microflora: principles and progress, Int J Hydrogen Energy, 2007, 32, 172–84. [42] Fan YT, Li CL, Lay JJ, Hou HW, Zhang GS. Optimization of initial substrate and pH levels for germination of sporing hydrogen-producing anaerobes in cow dung compost, Bioresour Technol, 2004, 91, 189–93. [43] de Vrije T, Budde MAW, Lips SJ, Bakker RR, Mars AE, Claassen PAM. Hydrogen production from carrot pulp by the extreme thermophiles Caldicellulosiruptor saccharolyticus and Thermotoga neapolitana, Int J Hydrogen Energy, 2010, 35, 13206–13. [44] Han W, Wang XN, Ye L, Huang JG, Tang JH, Li YF, Ren NQ. Fermentative hydrogen production using wheat flour hydrolysate by mixed culture, Int J Hydrogen Energy, 2015, 40, 4474–4480. [45] Panagiotopoulos IA, Bakker RR, de Vrije T, Koukios EG. Effect of pretreatment severity on the conversion of barley straw to fermentable substrates and the release of inhibitory compounds, Bioresour Technol, 2011, 102, 11204–11. [46] Sagnak R, Kargi F, Kapdan IK. Bio-hydrogen production from acid hydrolyzed waste ground wheat by dark fermentation, Int J Hydrogen Energy, 2011, 36, 12803–9. [47] Koutinas AA, Wang R, Webb C. Restructuring upstream bioprocessing: technological and economical aspects for production of a generic microbial feedstock from wheat, Biotechnol Bioeng, 2004, 85(5), 524–38. [48] Arifeen N, Kookos IK, Wang R, Koutinas AA, Webb C. Development of novel wheat biorefining: Effect of gluten extraction from wheat on bioethanol production, Biochem Eng J, 2009, 43, 113–21. [49] Zhang YZA, Sun Z, Leung CCJ, Han W, Lin SKC. Valorization of bakery waste for succinic acid production, Green Chem, 2012, 15, 690–5.

106 | 5 Utilization of food waste for fermentative hydrogen production

[50] Han W, Ye M, Zhu AJ, Huang JG, Zhao HT, Li YF. A combined bioprocess based on solid-state fermentation for dark fermentative hydrogen production from food waste, J Clean Prod, 2015, http://dx.doi.org/10.1016/j.jclepro.2015.08.072. [51] Han W, Lam WC, Melikoglu M, Wong MT, Leung HT, Ng CL, Yan P, Yeung SY, Lin SKC. Kinetic analysis of a crude enzyme extract produced via solid state fermentation of bakery waste, ACS Sustain Chem Eng, 2015, 3, 2043–2048. [52] Du CY, Lin SKC, Koutinas AA, Wang RH, Webb C. A wheat biorefining strategy based on solidstate fermentation for fermentative production of succinic acid, Bioresour Technol, 2008, 99, 8310–5. [53] Kountinas AA, Arifeen N, Wang R, Webb C. Cereal based biorefinery development: integrated enzyme production for cereal flour hydrolysis, Biotechnol Bioeng, 2007, 97, 61–72. [54] Ren NQ, Wang DY, Yang CP, Wang L, Xu JL, Li YF. Selection and isolation of hydrogen-producing fermentative bacteria with high yield and rate and its bioaugmentation process, Int J Hydrogen Energy, 2012, 35, 2877–82. [55] Wang R, Sharano SM, Gody LC, Melikoglu M, Webb C. Bioconversion of rapeseed meal for the production of a generic microbial feedstock, Enzyme Microb Technol, 2010, 47, 77–83. [56] Lee KS, Hsu YF, Lo YC, Lin PJ, Lin CY, Chang JS. Exploring optimal environmental factors for fermentative hydrogen production from starch using mixed anaerobic microflora, Int J Hydrogen Energy, 2008, 33, 1565–72. [57] Akutsu Y, Li YY, Tandukar M, Kubota K, Harada H. Effects of seed sludge on fermentative characteristics and microbial community structures in thermophilic hydrogen fermentation of starch, Int J Hydrogen Energy, 2008, 33, 6541–8. [58] Ren NQ, Wang BZ, Hung JL. Ethanol-type fermentation from carbohydrate in high rate acidogenic reactor, Biotechnol Bioeng, 1997, 54, 428–33. [59] Ren NQ, Cao GL, Guo WQ, Wang AJ, Zhu YH, Liu BF, Xu JF. Biological hydrogen production from corn stover by moderately thermophile Thermoanaerobacterium thermosaccharolyticum W16, Int J Hydrogen Energy, 2010, 35, 2708–12. [60] Wang J, Wan W. Effect of concentration on fermentative hydrogen production by mixed cultures, Int J Hydrogen Energy, 2008, 33(4), 1215–20. [61] Lay JJ, Fan KS, Chang J, Ku CH. Influence of chemical nature of organic wastes on their conversion to hydrogen by heat-shock digested sludge, Int J Hydrogen Energy, 2003, 28, 1361–7. [62] Sagnak R, Kargi F, Kapdan IK. Bio-hydrogen production from acid hydrolyzed waste ground wheat by dark fermentation, Int J Hydrogen Energy, 2011, 36, 12803–9. [63] Han W, Ye M, Zhu AJ, Zhao HT, Li YF. Batch dark fermentation from enzymatic hydrolyzed food waste for hydrogen production, Bioresour Technol, 2015, 191, 24–9.

Chao Chen and Tao Li

6 Bacterial dye-decolorizing peroxidases Biochemical properties and biotechnological opportunities 6.1 Introduction In biorefineries, processing biomass begins with separating lignin from cellulose and hemicellulose. The latter two are depolymerized to give monosaccharides (e.g. glucose and xylose), which can be converted to fuels or chemicals. In contrast, lignin presents a challenging target for further processing due to its inherent heterogeneity and recalcitrance. Therefore it has only been used in low-value applications. For example, lignin is burnt to recover energy in cellulosic ethanol production. Valorization of lignin is critical for biorefineries as it may generate high revenue. Lignin is the obvious candidate to provide renewable aromatic chemicals [1, 2]. As long as it can be depolymerized, the phenylpropane units can be converted into useful phenolic chemicals, which are currently derived from fossil fuels. In nature, lignin is efficiently depolymerized by rot fungi that secrete heme- and copper-containing oxidative enzymes [3]. Although lignin valorization is an important objective, industrial depolymerization by fungal enzymes would be difficult, largely due to difficulties in protein expression and genetic manipulation in fungi. In recent years, however, there is a growing interest in identifying ligninolytic bacteria that contain lignin-degrading enzymes. So far, several bacteria have been characterized to be lignin degraders [4, 5]. These bacteria, including actinobacteria and proteobacteria, have a unique class of dye-decolorizing peroxidases (DyPs, EC 1.11.1.19, PF04261) [6, 7]. These enzymes are equivalent to the fungal oxidases in lignin degradation, but they are much easier to manipulate as their functional expression does not involve post translational modification. Heme peroxidases play essential roles in biological activities. They are ubiquitous in all domains of life. These enzymes are structurally and functionally diverse although they all use heme in the catalytic cycle. Historically, these enzymes are classified into plant and animal peroxidase superfamilies based on primary sequence alignment and isolation origins. Later, phylogenic analysis revealed several groups of enzymes with distinct structural and catalytic properties [8, 9]. The resulting (super)families include peroxidase-catalase, peroxidase cyclooxygenase, peroxidasechlorite dismutase, di-heme peroxidase and haloperoxidase (Fig. 6.1). Only very recently, DyPs have been recognized as a novel class of heme proteins. They are structurally analogous to chlorite dismutases, a group of enzymes that are involved in chlorite detoxification [10]. However, their evolutional relationship is unclear. In contrast to other heme proteins, our knowledge about DyPs is very limited. However,

108 | 6 Bacterial dye-decolorizing peroxidases

their unique substrate specificity and catalytic property offer great opportunities for both exploring peroxidase enzymology, and biotechnological applications. The first Dyp-type protein was isolated in 1999 from Thanatephorus cucumeris Dec 1 (formerly Geotrichum candidum Dec 1, DyPDec1), a dye-decolorizing fungus [11]. The enzyme is a glycoprotein with low homology to all other known peroxidases. It has novel activity on anthraquinone dyes with low pH optimum (3.0–3.2) [12, 13]. More DyPs have been later found in prokaryotes. Although they all catalyze the oxidation of anthraquinone dyes, their primary sequence identity is low. Therefore, the DyPs are further classified into four classes from A to D (Fig. 6.1). The peroxidases in Classes A, B, and C are predominantly from bacteria, while those in Class D are largely from Basidiomycota. Many bacterial DyPs have shown to be attractive biocatalysts as they can oxidize a wide range of substrates such as synthetic dyes and lignin-derived chemicals [6, 7, 14–16]. Bacterial DyPs are a rich source of biocatalysts. For instance, a recent search of the Pfam database revealed 16 889 putative Dyp-type proteins, which are widely present in the 515 Bacteria species compared to the 38 Eukaryota species (Fig. 6.2). Despite the fact that increasing amount of DyP sequence information is available, only a handful of the peroxidases are well characterized. To realize their industrial potential, it is critical to understand their properties relevant to biocatalysis. In this chapter, we review the diverse biochemical properties of the bacterial DyPs that have been characterized. Their spectral properties and kinetic features are discussed on the basis of peroxidase mechanism. The physiological roles of the DyPs are discussed in the context of functional diversity. The structural characters of the DyPs are highlighted in relation to their underlying catalytic machinery. The biotechnological applications of peroxidases are reviewed to project potential use of bacterial DyPs as more efficient alternatives.

Haloperoxidase

Chlorite dismutase like proteins

Peroxidasecatalase

DyP class A DyP class B

Heme peroxidase

Peroxidasechlorite dismutase

Peroxidasecyclooxygenase

Dyp-type peroxidase DyP class C

Di-heme peroxidase

Chlorite dismutase

Fig. 6.1: Classification of heme peroxidase and Dyp-type peroxidase.

DyP class D

6.2 Biochemical properties |

109

s oe my pto e r St

Actinomycatales Actinobacteria

Actinobacteria ? c? Ba

Bac teri a

??

Root

e Ps

ia er ct ia er ba ct eo ba ot Pr eo t ro ap e m m ea ac Ga les ad da on na m mo o do as ud eu on Ps om ud

ceae xella Mora

acter etob Acin

e Ps

Enterobacteriales Enterobacteriaceae

acter Enterob

Fig. 6.2: Distribution of Dyp-type peroxidases across species. It is adapted from the Pfam database [17]. Bacteria proteins are colored as green and yellow, Eukaryota proteins are colored as purple, and unclassified proteins are colored as cyan (Search performed on September 15, 2015).

6.2 Biochemical properties Bacterial DyPs catalyze the oxidation of a wide range of substrates. The most notable ones are synthetic anthraquinones such as Reactive Blue series, which are poor substrates for other peroxidases. Similar to plant peroxidases, bacterial DyPs can also oxidize azo dyes, ABTS and small phenolic compounds including catechol, guaiacol, hydroquinone (HQ) and 2,6-dimethoxyphenol. Plant peroxidases are known to catalyze sulfide oxygenation. Similarly, an A-type TfuDyP from Thermobifida fusca was reported to convert aromatic sulfide to chiral sulfoxide products [18]. Comparing catalytic efficiency of peroxidases can be difficult since many reactions have different kinetic mechanisms, therefore entail different models [19]. For

110 | 6 Bacterial dye-decolorizing peroxidases

example, TcDyP from Thermomonospora curvata exhibits sigmodial kinetic profiles in the oxidation of ABTS and anthraquinone dyes [14], suggesting cooperation of multiple oxidation sites like fungal DyPs and versatile peroxidases [19–22]. Such a situation is simplified by fitting kinetic data to apparent saturation curves based on the app Michaelis–Menten equation. The resulting apparent Michaelis constant (Km ) and app the apparent turnover number (kcat ) are used to determine enzyme selectivity, the common parameters to compare different peroxidases (Tab. 6.1). Although DyPs from different classes have similar substrate range, fungal DyPs are generally more efficient than those from bacteria in the oxidation of ABTS and RB5. Only a few bacterial DyPs have comparable activity [14, 23]. OH HO

OH

O HO

O

OH

O OMe

OMe

OMe

MeO

OMe

OMe OR 1a: R=H 1b: R=Me

R

OMe

OMe 2

3

4a: R=H 4b: R=Me

Fig. 6.3: Structures of lignin model compounds.

Many bacterial DyPs from A–C classes can mediate the direct oxidation of lignin model compounds that requires high redox potential. TcDyP from T. curvata [14], DypB from Rhodococcus jostii RHA1 [6] and DyP2 from Amycolatopsis sp. 75iv2 [7] can oxidize guaiacyl glycerol-β-guaiacol ether (Fig. 6.3, 1a, ΔE = 0.6–0.8 V vs NHE) [24]. The degradation of 1a starts with Cα -Cβ bond cleavage, but the products can result from either degradation or polymerization, depending on the DyP and reaction conditions [6, 14]. Remarkably, some bacterial DyPs are active on nonphenolic lignin model compounds including veratryl alcohol (3, ΔE = 1.4 V vs NHE), guaiacyl glycerol-β-guaiacol ether (1b, ΔE > 1.0 V vs NHE), and 4-methoxymandelic acid (2, MMA, ΔE > 1.4 V vs NHE) [15, 18, 25]. Two Bacillus subtilis A-type DyPs sharing 96 % primary sequence identity have also showed promising activities. BsDyP from Bacillus subtilis KCTC2023 was found to catalyze the cleavage at Cα -Cβ of 1b. This is the first example of decomposing nonphenolic dimer by a bacterial DyP [15]. BsDyP from Bacillus subtilis 168 displayed peroxidase activities against syringaldehyde (4a) and acetosyringone (4b) [26]. TcDyP was found to oxidize MMA to give anisaldehyde as the product [13]. Moreover, A- and B-type DyPs have been demonstrated to decompose Kraft lignin without redox mediators [6, 16]. Notably, biochemical studies also have shown that temperature optimum for peroxidative activity is substrate dependent in some cases. For example, in BsDyP,

6.3 Physiological roles of bacterial DyPs

|

111

its maximal activities on lignin model compounds reached at 50 °C, while those for low redox potential substrates appeared at 30 °C. In nature, manganese peroxidase plays an important role in lignin degradation. It is a fungal enzyme that oxidizes Mn2+ to Mn3+ , which can diffuse in bulky lignin substrates to decompose the recalcitrant compounds. In the catalytic cycle, one equivalent H2 O2 oxidizes two equivalents of Mn2+ [27]. Clearly, manganese is a highly desirable mediator for oxidative degradation of lignin with bacterial DyPs. So far, manganese activities have been found in B- and C-type DyPs (Tab. 6.1). This activity can be critical for lignin degradation by these enzymes. For example, DypB from R. jostii RHA1 is activated by Mn2+ by 5–23 folds toward lignin model substrates. In wheat straw lignocellulose digestion, DypB is active only in the presence of 1 mM MnCl2 [6]. The catalytic efficiency of Mn2+ oxidation by the DypB is only 25.1 M–1 s–1 [28], approximately four orders of magnitude lower than that of bona fide manganese peroxidases [29, 30]. However, this efficiency is improvable by modifying the bacterial DyP. The substitution of Asn246 of DypB with Ala resulted in an 80-fold increase in the turnover of oxidation of Mn2+ to Mn3+ . Asn246 is the distal heme residue of DypB. This improvement is likely due to the higher reactivity of compound I, of which the half-life is shortened by three orders of magnitude by the mutation [31]. Two B-type DyP paralogs were found to be manganese active in Pseudomonas fluorescens Pf-5. One of the enzymes, DyP1B was found to degrade wheat straw lignocelluloses upon addition of Mn2+ . So far, the most app app manganese-reactive DyP is the C-type DyP2 from A. sp. 75iv2 [7], whose kcat /Km value 5 –1 –1 is 1.2 × 10 M s (Tab. 6.1), approaching the activity of fungal manganese peroxidases and versatile peroxidases [32, 33]. These examples strongly support that manganese can facilitate lignin degradation by bacterial DyPs as the redox mediator. In addition, several bacterial DyPs can catalyze a few extraordinary reactions. DyP2 catalyzed the conversion from MMA to anisaldehyde. This reaction is mediated by Mn2+ , and one additional equivalent of oxygen is required [7, 34]. An A-type DyP was proposed to catalyze deferrochelation, the removal of iron from heme for iron transportation and uptake [35]. Fungal DyPs also have shown oxygenase or hydrolase activities [36], however, these activities have not been found in hitherto characterized bacterial DyPs. Taken together, bacterial DyPs have been shown to be a group of catalytically versatile peroxidases.

6.3 Physiological roles of bacterial DyPs Based on their reactivity as well as their genomic context, bacterial DyPs have been proposed to fulfill various physiological roles. For example, a putative A-type DyP (VNG0798H) from Halobacterium salinarum has been suggested to be involved in regulatory machinery for oxidative stress. This protein was upregulated by reactive oxygen species in the native strain, and the survival rate for its knockout mutant was reduced by 99.9 % upon addition of 25 mM H2 O2 [41]. A study on E. coli gene expression profile

DyPDec1 AjP I AjP II TAP

D

ND: not detected;

DyP2 AnaPX

Pseudomonas aeruginosa PKE117 Shewanella oneidensis (strain MR-1) Pseudomonas putida MET94 Pseudomonas fluorescens Pf-5 Pseudomonas fluorescens Pf-5

DyPPa TyrA PpDyP DyP1BPl DyP2BPl

C

Rhodococcus jostii RHA1

DypB

B

Blank: not determined.

Thanatephorus cucumeris Dec 1 Auricularia auricula-judae Auricularia auricula-judae Termitomyces albuminosus

Amycolatopsis sp. 75iv2 Anabaena sp. strain PCC 7120

Thermomonospora curvata Thermobifida fusca Rhodococcus jostii RHA1 Bacillus subtilis KCTC2023 Pseudomonas fluorescens Pf-5

TcDyP TfuDyP DypA BsDyP DyPAPl

A

Source

proteins

Class

app

1.3 × 104 9.0 × 103 1.0 × 102

2.0 × 103 7.0 × 104 2.8 × 104 2.4 × 103

1.8 × 107 1.6 × 107 2.5 × 107

6.6 × 106

9.0 × 103 ND

6.5 × 105

1.7 × 107

8.0 × 103 1.2 × 104 5.8 × 103

RB4

ABTS

kcat /Km (M–1 s–1 )

app

Tab. 6.1: List of all biochemically characterized bacterial and fungal DyPs.

4.8 × 106 5.0 × 106 1.7 × 107

7.1 × 105 1.2 × 107

7.0 × 104 2.0 × 105

2.2 × 102

5.0 × 104

6.6 × 105

RB5 7.8 × 106 3.5 × 105

RB19

3.4 × 103 1.0 × 103 ND

3.2 × 102 ND

8.8 × 102

guaiacol

1.2 × 105

5.2 × 104 3.3 × 102 2.1 × 103

2.5 × 101

ND

ND

Mn2+

[11] [39] [39] [40]

[7] [23]

[37] [38] [26] [16] [16]

[28]

[14] [18] [28] [26] [16]

Ref.

112 | 6 Bacterial dye-decolorizing peroxidases

6.4 Catalytic mechanism of bacterial DyPs |

113

showed that the transcription of efeUOB (ycdNOB) operon is enhanced in acidic stress [42]. Later study suggested that EfeB (YcdB) and its paralog YfeX (B-type DyP) function are dechelatases for bacterial iron acquisition from heme, as their overexpression resulted in the accumulation of protoporphyrin IX (PPIX) [35]. This notion is supported by the genomic context, as efeB is downstream after efeU and efeO, which encode a high-affinity iron transporter and an iron uptake component, respectively. However, purified EfeB does not catalyze iron removal from hemin, suggesting that other mechanisms might be involved in the deferrochelation activity [43]. The function of YfeX as a deferrochlatase is also controversial. It has been proposed to be a porphyrinogen oxidase based on its activity on protoporphyrinogen IX and coproporphyrinogen III [44]. The location of an enzyme also provides clues to its physiological role. A-type DyPs are predicted as periplasmic proteins due to their unique N-terminal twinarginine translocation (TAT) signal peptides for translocation. So they may fulfill different roles from the DyPs in other classes. Still, this rule is not general as only some A-type DyPs have been experimentally confirmed as TAT substrates [18, 45]. DypB, the B-type DyP from R. jostii RHA1, has been shown to oxidize polymeric lignin and lignin model compounds. Also, deletion of dypB gene in R. jostii RHA1 reduced lignin degradation activity [6]. Still, other evidence suggested that the physiological role of DypB may not be associated with lignin degradation [46]. For example, R. jostii RHA1 cannot grow on lignin as sole carbon source. The gene dypB homologs are found to be co-localized and probably co-expressed with enc in many bacteria including R. jostii RHA1 [47–49]. The gene enc encodes encapsulin, a bacterial icosahedral nanocompartment, which can encapsulate functional proteins such as ferritin. Therefore, the role of DypB remains to be clarified. C-type DyPs are more efficient catalysts than those in A- and B-type. They displayed comparable manganese peroxidase activity to those of plant peroxidases [7]. The biochemical properties suggest that C-type DyPs are more likely to function as lignin-degrading peroxidases. However, their genes are organized in highly diverse genomic contexts, suggesting many different roles in biological processes. The identification of natural substrates for these enzymes will be critical to unveil their physiological roles.

6.4 Catalytic mechanism of bacterial DyPs Bacterial DyPs have been proposed to employ the mechanism similar to that of classical peroxidases [13, 36]. The catalytic process typically requires two single-electron oxidation. In the first step, a resting ferric enzyme reacts with hydrogen peroxide to yield a high-valent [Fe4+ =O]+• that is designated as compound I; the oxoferryl species is then reduced by one equivalent electron from a reducing substrate to form [Fe4+ =O]+ intermediate that is designated as compound II; and last, compound II undergoes a second reduction by one equivalent electron to restore the [Fe3+ ] resting state of the

114 | 6 Bacterial dye-decolorizing peroxidases

peroxidase. Since these peroxidative reactions are generally irreversible, this mechanism is referred as peroxidase ping-pong kinetics (Fig. 6.4) [50]. Although the steady state data for a typical peroxidase still fit the Michaelis–Menten equation, the resulting parameters have entirely different interpretations. For example, the value of Km for substrate is typically k1 [H2 O2 ]/k3 , where k1 and k3 represent reaction constants for compound I formation and compound II reduction, respectively (Fig. 6.4). H2O2

TcDyP–0

H2O

k1

AH2

TcDyP–I

AH

k2

AH2

TcDyP–II

AH + H2O

k3

TcDyP–0

Fig. 6.4: The proposed peroxidase ping-pong kinetic mechanism in TcDyP. k1 , k2 , and k3 represent rate constants for compound I formation, compound II formation and its reduction.

Soret band Q bands

0.30

0.030 0.025

0.25 Absorbance

0.020

0.20

0.015 CT bands 0.010

0.15

0.005 0.000

0.10

500

600

700

600

700

0.05 0.00 400

500 Wavelength (nm)

Fig. 6.5: Spectral characteristics of wt-TcDyP, illustrating Soret, Q and charger transfer (CT) bands.

The transition of the π-electrons in peroxidase porphyrin generates unique UV-Vis spectrum. Like other heme containing peroxidases, the electronic adsorption spectra of DyPs have Soret band, Q (α and β) and charge transfer (CT) bands as shown in Fig. 6.5. The change of microenvironment of porphyrin usually causes the shift of these bands. This change offers important opportunities to study the mechanism in peroxidase catalytic cycle, including characterizing the reaction intermediates, measuring the rate of their formation and transformation, and the significance of amino acids in the active sites.

6.4 Catalytic mechanism of bacterial DyPs |

115

Tab. 6.2: Absorption maxima of resting state and oxidized intermediates of heme peroxidases (adopted from [14]). Name

pH value

Resting state

Compound I

Compound II

Ref.

wt-DypA

7.5

408, 502, 632

ND

419, 528, 557, 619

[28]

wt-DypB

7.5

404, 503, 634

400, 580, 613, 648

ND

[28]

TcDyP

3.0

406, 509, 542(sh), 573, 630

406, 510, 547, 569, 621, 646(sh)

410, 515, 549, 621, 662 (sh)

[14]

7.8

406, 508, 544(sh), 567(sh), 624

407, 512, 546, 623, 648 (sh)

416, 530, 555, 623, 666

DyPDec1

3.2

406, 506, 636

401, 530, 556, 615, 644(sh)

399, 529, 555, 615, 644(sh)

[13]

HRP

6.6

403, 498, 640

400, 525(sh), 577, 622(sh), 651

420, 527, 555

[57]

LiP

6.0

408, 496, 630

408, 550, 608, 650

420, 525, 556

[58]

MnP

4.5

406, 502, 632

407, 558, 617, 650

420, 528, 555

[59]

The sh and ND mean a shoulder peak and not detectable, respectively.

In theory, adding H2 O2 to resting state DyP causes the formation of compound I. The formation of compound II from compound I is carried out by adding one equivalent of one-electron reductant. The addition of second equivalent reductant can convert compound II back to the resting state. Compound I and compound II can be distinguished by electron paramagnetic resonance (EPR) since it has an unpaired electron. So far, only a few bacterial DyPs have been studied for reactive intermediates with UV-Vis (Tab. 6.2). In reality, however, the mechanism is much more complicated. Upon reacting with one equivalent of H2 O2, the D-type DyP from T. cucumeris Dec 1 underwent intensity change of Soret peak and emergence of new peaks in Q bands and CT band in 0.2 min, suggesting the formation of compound I. Adding one more equivalent H2 O2 did not cause additional changes. However, the enzyme returned to resting state after 20 min, suggesting that compound I is reduced [13]. DypA and DypB are two peroxidases in R. jostii RHA1 with different substrate specificities: The A-type DypA app app prefers Reactive Blue 4 (kcat /Km = 12800 ± 600 M−1 s−1 ), while the B-type DypB app app app app prefers ABTS (kcat /Km = 2000 ± 100 M−1 s−1 ) and also oxidizes Mn2+ (kcat /Km = −1 −1 25.1 ± 0.1 M s ). When reacting with H2 O2 , DypA was converted into an intermediate with features of compound II, while DypB slowly (second-order rate constant ~ 1.80 × 105 M–1 s–1 ) turned into a compound I-like protein-based radical species [28]. TcDyP, the A-type peroxidase from T. curvata is more systematically studied with UV-Vis spectroscopy [14]. The spectral transition was measured for the reaction of TcDyP with H2 O2 . Fitting multiwavelengthdata revealed the formation of two intermediates: one is compound I (TcDyP-I), and the other is proposed to be a compound II-like

116 | 6 Bacterial dye-decolorizing peroxidases 408

406

416

416

0.20

524 506 555

0.02

0.2

0.01 0.1 450

500

450

500

550

600

511

0.00 700

650

400

550

600

650

0.15

0.02

0.10

0.01

450 0.00 350

700

400

450

(b)

Wavelength (nm)

(a)

0.03 555

0.05

0.0 350

524

0.03

Absorbance

Absorbance

0.3

500

500

550

600

650

0.00 700

550

600

650

700

Wavelength (nm) 408 412

406

0.6

512 546

0.30

0.04

0.02 509 0.01

0.2

450

500

550

500

550

600 650

0.20

0.04

0.15 0.02 0.10 0.05

0.00 700

0.0

450

500

550

600

650

0.00 700

450

500

550

600

650

700

0.00 350

(c)

Absorbance

Absorbance

0.03

410

0.4

0.06

551 526

0.25

400

450

600

Wavelength (nm)

650

350

700

(d)

400

Wavelength (nm)

Fig. 6.6: Spectral changes in wt-TcDyP and TcDyP-H281A catalytic cycle. A: Reaction of wt-TcDyP with H2 O2 . B: Reaction of TcDyP-I with hydroquinone in a sequential mixing mode. C: Reaction of TcDyP-II with hydroquinone in a sequential mixing mode. D: Reaction of TcDyP-H281A with H2 O2 . TcDyP-0, TcDyP-I, and TcDyP-II (TcDyP-II like) compounds are shown in blue, green and red curves, respectively. Arrows indicate changes of absorbance over time. This research was originally published in the Journal of Biological Chemistry [51].

decay product (TcDyP-II like) (Fig. 6.6 (a)). In the formation of TcDyP-I, there is a decrease of the Soret band absorption at 406 nm. By using this change, the second order rate constant was determined to be 5.92 × 106 M–1 s–1 at pH 7.8. At pH 3.0, this rate was marginally reduced to 4.06 × 106 M–1 s–1 . Although this trend is similar to that of DypB [28], TcDyP-I formation is about 30 fold faster than that of horseradish peroxidase (1.7 × 106 M–1 s–1 ) [52]. Compound II-like decay product of TcDyP could be a protein radical that has been observed in many heme peroxidases [53, 54]. Intriguingly, the transient intermediates spontaneously returned to the resting state, a phenomenon that has been reported in other DyPs and peroxidases [13, 55]. The formation of compound II of TcDyP was achieved by using one equivalent hydroquinone (HQ) to reduce TcDyP I. A second equivalent of HQ converted TcDyP-II back to resting state. The rate for TcDyP-II formation was determined at 416 nm (TcDyP-II Soret band, pH7.8), while

6.5 Structure-function relationship in bacterial DyPs |

117

O Fe3+ TcDyP–0

OH

6. 3

6x

10 3

O

M –1 s –1 (pH 3

.0)

.8) H 7 .0) –1 (p 3 –1 s 1 pH 6 M 1 – ( 10 6 M– s 2x 0 5.9 6 x 1 4.0

OH

H2O2

H2O

.+

+ 2.24 x 104 M–1s–1(pH 7.8)

O Fe4+

O Fe4+

TcDyP–II

TcDyP–I O

O

HO

OH Decay TcDyP–II–like compound

Fig. 6.7: Catalytic cycle of wt-TcDyP. Second-order rate constants are shown in each step [51].

that for TcDyP-II reduction was measured at 406 nm (TcDyP-0 Soret band) (Fig. 6.7). The reduction of TcDyP-II to regenerate TcDyP-0 turned out to be the slowest step in HQ-oxidation. In the full catalytic cycle of TcDyP, compound II reduction is the ratelimiting step. It is noteworthy to mention that UV-Vis spectroscopy has its limitation to study peroxidase mechanism when the Soret band overlaps with the spectrum of a dye substrate such as ABTS. This issue can be mitigated by incorporating additional reaction to eliminate absorption change due to ABTS oxidation [56].

6.5 Structure-function relationship in bacterial DyPs Bacterial DyPs are mainly assembled in dimers, while fungal DyPs are exclusively monomers. Sequence analysis indicates that fungal DyPs have numerous C-terminal insertion sequences, which are postulated to prevent the enzymes from dimerization [60]. In bacterial DyPs, the dimerization is usually caused by hydrophobic interaction between monomers. Higher order oligomerization has also been observed (Tab 6.3). The dimer of BtDyP from Bacteroides thetaiotaomicron VPI-5482 further associates to form a hexamer, mainly via hydrophobic interaction [58]. Nevertheless, there are exceptions to the dimeric organization. TfuDyP from T. fusca, an A-type DyP, is active as a monomer, suggesting that dimerization may not involve the DyP activity [18]. DyP2 is a C-type DyP from Amycolatopsis. Dynamic light scattering measurement suggested

118 | 6 Bacterial dye-decolorizing peroxidases Tab. 6.3: List of all available structures of bacterial DyPs. Proteins

Organisms

Class

PDB code

Uniprot ID

Quaternary structures

EfeB

Escherichia coli

A

2Y4F (heme); 3O72 (heme)

Q8XAS4

Dimer

DyP

Thermobifida cellulosilytica

A

4GS1 (heme)

U3KRF5

Dimer

SCO3963

Streptomyces coelicolor

A

4GT2 (heme)

Q9ZBW9

Dimer

SCO2276

Streptomyces coelicolor

A

4GRC (heme)

Q9RKQ2

Dimer

DypB

Rhodococcus jostii RHA1

B

3QNS (heme); DypB N246A: 4HOV (heme, Mn)

Q0SE24

Hexamer

TyrA

Shewanella oneidensis (strain MR-1)

B

2IIZ (heme); 2HAG (apo)

Q8EIU4

Dimer

BtDyP

Bacteroides thetaiotaomicron VPI-5482

B

2GVK

Q8A8E8

Hexamer

SCO7193

Streptomyces coelicolor

B

4GU7 (heme)

Q9FBY9

Dimer

DyP2

Amycolatopsis sp. 75iv2

C

4G2C (heme, Mn)

K7N5M8

Monomer (crystal); Dimer (PISA predicted)

that the enzyme existed as oligomer in solution. In crystallization, this enzyme existed as a dimer with no strong interaction at the interface. The physiological organization of DyP2 is inconclusive due to the discrepancy between the experiments [7]. DyPs are very different from classical peroxidases, which are primarily α-helical proteins [61]. All DyPs share an α + β ferredoxin-like fold that resembles the tertiary structure of chlorite dismutases (Clds) [61]. Therefore, DyPs and Clds (like) proteins have been proposed to constitute a new superfamily even though they have no significant sequence identity [10]. The bacterial DyP protomer is divided into two ferredoxin-like domains: an N-terminal domain and a larger C-terminal domain. Each domain consists of a four-stranded antiparallel β-sheet surrounded by α-helices. The conserved topology in each domain may reflect the result of gene duplication. In accordance with the 1 : 1 stoichiometric ratio, each subunit has one heme B binding to the C-terminal domain. Structure-based pairwise alignment of DyPs from different types revealed that they share relatively low identity (Dali Z-score < 32, root mean square deviation > 2.5 Å). However, critical residues in β-sheet cores (heme binding pocket) are highly conserved (Fig. 6.8) [62]. At the reactive site, the most critical residues are the axial ligands for the heme. In all DyPs, the heme iron is coordinated to the Nε atom of a proximal histidine in

6.5 Structure-function relationship in bacterial DyPs |

(a)

(b)

D235

R347

(c)

D153

F368

119

R244

F261

D192

R348

F378

H323 H329

H226

(d)

D414

D288

E389 (e)

(f)

Fig. 6.8: Overall structure and heme environment in bacterial DyPs. (a), (b), and (c) represent overall structures of A-type EfeB (PDB code 3O72), B-type DypB (PDB code 3QNS) and C-type DyP2 (PDB code 4G2C), respectively. (d), (e), and (f) represent their respective active sites including heme, its surrounding conversed residues and oxygen species, which are shown in a stick model (green). Structural elements including α-helices (cyan), β-sheet (magenta) as well as loops (salmon) are shown in a cartoon model.

C-terminal domain, and the Nδ atom of the proximal ligand is in hydrogen bonding distance to the carboxylate of an acidic residue (Asp or Glu). It is very reminiscent of a widespread motif of Fe-His-Asp triad in metalloenzymes [63]. The interaction between aspartic acid and histidine has been proved to be critical to the push and pull effect involved in compound I formation in several peroxidases [64]. Therefore, replacing the proximal histidine is expected to reduce the peroxidase activity due to ligand loss. Site-directed mutagenesis analysis in TfuDyP supported this proposition: the substitution of histidine with alanine abolished heme binding, and the distinctive Soret band disappeared [18]. Similar mutation in TcDyP also caused the loss of activity, but detailed study revealed different mechanism. Based on transient kinetics, the H312A mutant was able to form compound I and II with the activities comparable to those found in the wild-type enzymes. However, the mutant cannot be reduced from compound II to the resting state with HQ. Hence, H312A cannot complete a catalytic

120 | 6 Bacterial dye-decolorizing peroxidases

cycle. The wild-type TcDyP and the H312A mutant have very similar spectral features for all intermediates (TcDyP-0, TcDyP-I, and TcDyP-II) (Fig. 6.6 (d)), suggesting that the function of H312A was partially rescued by the other residues in the region [14]. Indeed, the H312 is located on a short α-helix between two large loops (G276–A311 and H317–P329) in the homology model (Fig. 6.9) [14]. The high structural flexibility in this region may allow other residues to fill the role of the proximal histidine in H312A to carry out the catalysis to the stage of compound II.

Loop 200–229 Loop 317–329

Loop 317–329

Loop 317– 329

Loop 200–229

Propionate entry excess

Loop 276–311

(a)

90°

Loop 200– 229

Loop 276–311

Loop 276–311

(b)

Fig. 6.9: The TcDyP model structure. (a): Ribbon presentation of the TcDyP model including a loop of 200–229 (red) connecting the two domains, and two loops of 276–311(yellow) and 317–329 (magenta) flanking the short α-helix where the proposed axial H312 is situated. The heme and the proposed catalytic residues are colored in green and cyan, respectively. (b): Surface presentation of the TcDyP model with two orientations (related by 90° rotation) and the three loops.

On the distal face of the heme, bacterial DyPs have three highly conserved residues including an aspartate, an arginine and a phenylalanine (Fig. 6.8 (c), (d), and (e)). For DyPDec1 from T. cucumeris Dec1, the mutant D171N lost reactivity with H2 O2 as indicated by UV-Vis spectroscopy [13]. The conversed aspartate residue is proposed to function as an acid-base catalyst by accepting a proton from the peroxide prior to forming compound I [13]. This role is equivalent to that of the distal histidine in cytochrome c peroxidase [63] or that of glutamate in chloroperoxidase [65]. This notion is also supported by the fact that all bacterial DyPs have pH optima around the pKa of the carboxylate of the aspartic acid [8]. However, the mutation study with DypB from R. jostii RHA1 suggested that the conserved aspartate is not essential for peroxidase activity. The D153A mutant can catalyze the formation of compound I just as efficiently as the wild-type enzyme, although the stability of compound I was dramatically reduced [66]. Indeed, the aspartate residue is most likely to play divergent roles in DyPs. The role of the conserved arginine is also under debate. In DyPDec1, the guanidine of R329 is proposed to stabilize the negatively charged peroxide during O–O

6.5 Structure-function relationship in bacterial DyPs |

121

fission [13]. For DypB from R. jostii RHA1, the R244L mutation prevented the formation of compound I, indicating that the conserved arginine plays an essential role in catalysis [66]. The position of the conserved arginine suggests additional functions. Based on the available DyP structures, the arginine is the closest residue to the heme iron (~ 4.3 Å), and forms a hydrogen bond with the propionate moiety on pyrrole D in a bent conformation (Fig. 6.8 (d), (e), and (f)). This particular conformation suggests that the arginine helps to position the heme in the active site. Spectroscopic study of DyPs in solution shows that resting state DyPs contain high-spin, penta-coordinated hemes. In crystal structure, however, the sixth coordination site on the distal face is occupied by varying species. The DypB heme had an iron-coordinated solvent species [28], while the heme iron in EfeB bounded to an oxygen molecule [43]. Their presence may not represent the native state as protein crystallization and X-ray structure determination may create artifacts [67]. It is unclear how heme is incorporated into the deeply buried cavity in DyPs. Structural alignment of apo-TyrA (PDB code 2HAG) and holo-TyrA (PDB code 2IIZ) revealed little change upon heme binding except side chain rotation in several residues [38], suggesting that other factors are likely involved in heme insertion. It has been found that the heme binding pocket in DyPs is usually surrounded by a long flexible loop that bridges the N- and C-terminal domains. In EfeB, the loop include residue 222–243 [43], while in TcDyP, the equivalent loop include residue 200–229, both at the confluence of the two domains (Fig. 6.9). The flexibility of this loop comes from the relative movement of the two domains, and is speculated to contribute to not only heme incorporation, but also the catalytic turnover [43]. In EfeB, the heme is situated at the end of a propionate channel on the proximal side [43]. The name comes from the fact that the two propionic acid moieties of heme adopt outward orientation. This channel is proposed to be a common substrate access route for small substrates, as it is observed in all characterized DyPs (Fig. 6.9). Some DyPs also have an additional distal channel, but it is relatively narrow, therefore can only accommodate even smaller substrates. In handling bulky substrates such as lignin polymer or dyes, another mechanism is needed. Structural study provides insight to the mechanism of Mn2+ oxidation. Manganese plays a critical role in lignin degradation as a diffusible mediator that can access the interior of the polymer. Kinetic study shows that B- and C-type DyPs can employ Mn2+ /Mn3+ interconversion to mediate lignin oxidation. In DypB-N246A, the bound manganese is 4.2 Å away from the heme, a distance similar to those found in fungal manganese peroxidases [68]. It is probably oxidized in the Fe-heme site, although Mn2+ is directly coordinated by solvent species instead of the typical acidic residues [31]. By contrast, in DyP2, the Mn2+ cannot be oxidized directly by the iron as it is held by carboxylates of three glutamic acid residues that are ~ 16 Å away from the heme [7]. A long distance electron transfer (LRET) mechanism has therefore been proposed for DyP2, since a midway Tyr188 can facilitate electron relay as suggested by the mechanism in a di-heme peroxidase [69].

122 | 6 Bacterial dye-decolorizing peroxidases

A LRET mechanism may also play an important role in DyP catalysis for bulky substrates such as lignin or synthetic dyes, since they are occluded from the deep buried heme center. For Kraft lignin degradation, structural analysis suggests that DypB may employ a radical relay network between the protein surface and the active site [28]. Stronger evidence comes from the systematic study of the DyP from Auricularia auricula-judae, a common fungus that grows on wood. In the oxidation of Reactive Blue 19, the enzyme mainly employs Trp377 as the oxidation site at the beginning of an LRET pathway. The turnover involving the LRET mechanism showed a turnover of 200 s–1 , proving its importance over the direct heme access route, which showed a turnover of 20 s–1 [20].

6.6 Biotechnological opportunities Bacterial dye-decolorizing peroxidases emerge as a promising biocatalyst by offering many advantages over fungal DyPs and classical peroxidases. Advances in nextgeneration sequencing and metagenome provide a vast variety of bacterial DyPs that potentially have novel activities. Many characterized enzymes have already shown efficient activities toward an array of substrates, including recalcitrant dyes and lignin model compounds. They also display maximal activities under acidic conditions, permitting facile integration with lignocellulose hydrolysis under acidic conditions. As prokaryotic proteins, they are easier to optimize via protein engineering, and can be synthesized in high yield in standard E. coli expression systems [70–72]. Taken together, bacterial DyPs are attractive tools for biotechnological applications.

6.6.1 Lignin valorization and fine chemicals Lignin in cell walls is the second most abundant biopolymer after cellulose which comprises 25 % of plant biomass on average. Large quantities of lignin are generated as a byproduct from pulp and paper manufacture via the Kraft process. Potentially even larger amounts of lignin will be produced in the biofuel industry. It has been estimated that by 2022 the US bioethanol industry will generate about 60 million tons of lignin annually [1]. Power production from lignin combustion is one of the easiest ways to use lignin byproduct, but the value of energy from this approach is very low. Lignin is expected to become an alternative feedstock for aromatic chemicals, most of which are currently derived from the BTX process in the petroleum industry. Industrial biotechnology is expected to make great contributions to lignin valorization, which involves lignin depolymerization into small molecules and transforming them into value-added products [73]. In nature, lignin is degraded by extracellular lignin or Mn peroxidases to give a plethora of aromatic molecules [74]. These aromatics include β-Aryl ethers, biphenyls, diaryl propanes, pinoresinols, phenyl-

6.6 Biotechnological opportunities

|

123

coumaranes, and ferulic acid, etc. They are mediated by their dedicated pathways to give mono-aryl compounds. These compounds are degraded by the β-ketoadipate pathway, via either protocatechuate branch or catechol branches, to give β-ketoadipic acid as the common intermediate before joining the central metabolism i.e. the TCA cycle [75, 76]. Industrial biotechnology allows integration of lignin depolymerization with further transformation, thereby permitting direct conversion of lignin into useful products. This is a distinctive advantage over chemical processes, in which lignin needs to be depolymerized into process compatible monomers. In the Gram-positive soil bacterium R. jostii RHA1, knocking out the vanillin dehydrogenase gene enabled vanillin production directly from wheat straw [77]. The peroxidases were not only responsible for lignin degradation, but also produced the vanillin product from β-aryl ethers via the peroxidation reaction [6]. The mutant strain produced up to 96 mg/l vanillin after six days when grown on minimal medium supplemented with 2.5 % wheat straw lignocelluloses. In a different strain of R. jostii RHA1, aromatic dicarboxylic acids can be produced directly from lignin by introducing exogenous genes of protocatechuate 4,5dioxygenase (LigAB) or protocatechuate 2,3-dioxygenase (PraA) to work with DypA and DypB in aryl cleavage [78]. Lignin depolymerization gives a heterogeneous mixture, thus presenting a barrier to producing homogeneous product via biotransformation. This problem can be overcome by a biological funneling strategy, in which the diverse aromatics are catabolized to give a common intermediate, which can be used by the same organism to produce useful products. Gram-negative bacterium P. putida KT2440 is a natural aromatic-catabolizing organism with a diverse metabolic repertoire which can transform diverse aromatics from lignin depolymerization to join the β-ketoadipate pathway. It showed the ability to produce medium chain length polyhydroxyalkanoates (PHA) from pretreated lignin enriched biomass [79]. To enhance muconate production, this strain was improved by genetic manipulation involving deletion of genes encoding PcaHG and CatBC and insertion of genes encoding AroY and DmpKLMNOP [80]. The engineered strain could use lignin as the substrate, and reached the highest muconate yield of 13.5 g/l in fed-batch mode using p-coumarate as a carbon source.

6.6.2 Dye decolorization in wastewater treatment Synthetic dyes are chemically stable xenobiotics that are used in the manufacture of textiles, cosmetics, food and pharmaceuticals. The dye-contaminated wastewater needs to be treated before its discharge into the environment. Every year over 105 tons of dyes were released into wastewater stream [81], of which azo and anthraquinonic dyes are the major components. Treating wastewater presents a great opportunity for biological decolorization.

124 | 6 Bacterial dye-decolorizing peroxidases

Bacterial DyPs have shown up to 40-fold higher decolorizing activities than other dye-degrading enzymes including laccase and azoreductase [26]. In the oxidation of a recalcitrant dye RB5, the catalytic efficiency reached as high as 1.2 × 107 M–1 s–1 . Some DyPs have broad substrate specificities. For example, PpDyP from P. putida MET 94 and BsDyP from B. subtilis have similar activities towards azo and anthraquinonic dyes [26]. The degradation pathway has only been studied for RB5 degradation [36]. Analysis of the peroxidation with DyPDec1 showed that the products from RB5 included three major products, including two red-brown compounds and phthalic acid. Adding TcVP1, a versatile peroxidase from the same organism further rendered red-brown products colorless [82]. These results suggest that the decolorization of RB5 may involve two consecutive enzyme reactions in T. cucumeris Dec 1. There are only a limited number of known enzymes that can efficiently decolorize industrial dyes [83]. Discovery of new DyPs, and integration of multiple enzyme reactions in one organism should provide many options for dye decolorization and detoxification [84].

6.6.3 Other potential industrial applications The exploration of novel antimicrobial therapeutics is critical as a public health crisis has often arisen from issues surrounding antibiotic-resistant pathogens. A-, B-, and Ctype DyPs are predominantly found in bacteria. They are absent in higher eukaryotic organisms. For very few known bacterial DyPs, their role can be unique and critical. EfeB plays a key role in iron acquisition as a deferrochelase in E. coli [42]. It may serve as a target for antimicrobial drug development [43]. In the oxidation of peroxidase by hydrogen peroxide, the oxidation state of Fe is changed from III to IV. Direct reduction of the oxidized Fe(IV) to Fe(III) with electric current provide the basis for amperometric determination of H2 O2 concentration [85]. In sensor development, the peroxidase is immobilized on the electrode, so that the signal transduction takes place directly between the enzyme and the electrode. Historically, horseradish peroxidase is the most popular enzyme for H2 O2 sensor development because it is a well-studied enzyme. As alternatives to horseradish peroxidase, bacterial DyPs offer all the advantage as prokaryotic enzymes. Recently, B-type PpDyP was immobilized on Ag electrodes, and was found to have efficient electro-catalytic activity towards the peroxide [86]. Raman spectroscopic analysis revealed that the immobilized PpDyP preserved structural integrity of heme environment, which however was often destabilized in other enzymes upon immobilization [87, 88]. This result should encourage more work to explore the advantages that DyPs may offer.

6.7 Conclusions and perspectives |

125

6.7 Conclusions and perspectives Bacterial DyPs are novel heme peroxidases with highly diversified substrate specificities and physiological roles. Although they are structurally distinct from classical peroxidases, they share the same mechanism that involves the typical intermediates at different oxidation stages. Kinetic study contributes to the understanding of catalytic mechanism, substrate selectivity, and roles of amino acid residues in enzyme catalysis. In the active site of DyPs, the conserved amino acid residues at the distal side of heme are unique, including an aspartate, an arginine, and a phenylalanine. Their roles in catalysis have been speculated to be acid/base catalysis, charge stabilization, or even heme positioning based on the study in a few bacterial DyPs. The Mn activity found in B- and C-type DyPs is significant for their activity on lignin degradation. The mechanism, which may involve long range electron transfer, is unique and important for the oxidation of bulky substrates. Therefore it deserves more comprehensive study. Although the diversity of bacterial DyPs is well recognized, only a limited number of them are well characterized in terms of biochemical property. The physiological roles of these DyPs are not necessarily related to their peroxidase activity as shown by their biochemical property and genomic context. Increasing numbers of DyPs have been found in the search for biocatalysts in industrial applications including synthetic dye degradation or lignin valorization. Finding the right biocatalysts depends on the diversity in the pool of bacterial DyPs. Many more unique DyPs are expected to be found in metagenome sequences from diverse environments. For process development, the operational performance of a peroxidase is expected to be improved by in vitro evolution. Together, these activities should enable the practical application with bacterial DyPs.

Disclaimer This work is not a product of the United States Government or the United States Environmental Protection Agency, and the authors are not doing this work in any governmental capacity. The views expressed are those of the authors only and do not necessarily represent those of the United States or the US EPA.

References [1]

[2]

Holladay JE, White JF, Bozell JJ, Johnson D. Top Value Added Chemicals from Biomass. Volume II: Results of Screening for Potential Candidates from Biorefinery Lignin. US Department of Energy, 2007. Ragauskas AJ, Beckham GT, Biddy MJ et al. Lignin valorization: improving lignin processing in the biorefinery, Science, 2014, 344, 1246843.

126 | 6 Bacterial dye-decolorizing peroxidases

[3]

[4] [5]

[6] [7] [8] [9] [10]

[11] [12]

[13]

[14]

[15]

[16]

[17] [18]

[19]

[20]

Martinez AT, Speranza M, Ruiz-Duenas FJ et al. Biodegradation of lignocellulosics: microbial, chemical, and enzymatic aspects of the fungal attack of lignin, Int Microbiol, 2005, 8, 195–204. Bugg TD, Ahmad M, Hardiman EM, Singh R. The emerging role for bacteria in lignin degradation and bio-product formation, Curr Opin Biotechnol, 2011, 22, 394–400. Brown ME, Walker MC, Nakashige TG, Iavarone AT, Chang MC. Discovery and characterization of heme enzymes from unsequenced bacteria: application to microbial lignin degradation, J Am Chem Soc, 2011, 133, 18006–9. Ahmad M, Roberts JN, Hardiman EM, Singh R, Eltis LD, Bugg TD. Identification of DypB from Rhodococcus jostii RHA1 as a lignin peroxidase, Biochemistry, 2011, 50, 5096–107. Brown ME, Barros T, Chang MC. Identification and characterization of a multifunctional dye peroxidase from a lignin-reactive bacterium, ACS Chem Biol, 2012, 7, 2074–81. Zamocky M, Hofbauer S, Schaffner I et al. Independent evolution of four heme peroxidase superfamilies, Arch Biochem Biophys, 2015, 574, 108–19. Torres E, Ayala M. Biocatalysis based on heme peroxidases: peroxidases as potential industrial biocatalysts. New York: Springer-Verlag, 2010. Goblirsch B, Kurker RC, Streit BR, Wilmot CM, DuBois JL. Chlorite dismutases, DyPs, and EfeB: 3 microbial heme enzyme families comprise the CDE structural superfamily, J Mol Biol, 2011, 408, 379–98. Kim SJ, Shoda M. Purification and characterization of a novel peroxidase from Geotrichum candidum dec 1 involved in decolorization of dyes, Appl Environ Microbiol, 1999, 65, 1029–35. Sugano Y, Nakano R, Sasaki K, Shoda M. Efficient heterologous expression in Aspergillus oryzae of a unique dye-decolorizing peroxidase, DyP, of Geotrichum candidum Dec 1, Appl Environ Microbiol, 2000, 66, 1754–8. Sugano Y, Muramatsu R, Ichiyanagi A, Sato T, Shoda M. DyP, a unique dye-decolorizing peroxidase, represents a novel heme peroxidase family: ASP171 replaces the distal histidine of classical peroxidases, J Biol Chem, 2007, 282, 36652–8. Chen C, Shrestha R, Jia KM et al. Characterization of dye-decolorizing peroxidase (DyP) from Thermomonospora curvata reveals unique catalytic properties of A-type DyPs, J Biol Chem, 2015, 290, 23447–63. Min K, Gong G, Woo HM, Kim Y, Um Y. A dye-decolorizing peroxidase from Bacillus subtilis exhibiting substrate–dependent optimum temperature for dyes and beta-ether lignin dimer, Sci Rep, 2015, 5, 8245. Rahmanpour R, Bugg TD. Characterisation of Dyp-type peroxidases from Pseudomonas fluorescens Pf-5: Oxidation of Mn(II) and polymeric lignin by Dyp1B, Arch Biochem Biophys, 2015, 574, 93–8. Finn RD, Bateman A, Clements J et al. Pfam: the protein families database, Nucleic Acids Res, 2014, 42, D222–30. van Bloois E, Torres Pazmino DE, Winter RT, Fraaije MW. A robust and extracellular hemecontaining peroxidase from Thermobifida fusca as prototype of a bacterial peroxidase superfamily, Appl Microbiol Biotechnol, 2010, 86, 1419–30. Strittmatter E, Serrer K, Liers C et al. The toolbox of Auricularia auricula-judae dye-decolorizing peroxidase – Identification of three new potential substrate-interaction sites, Arch Biochem Biophys, 2015, 574, 75–85. Linde D, Pogni R, Canellas M et al. Catalytic surface radical in dye-decolorizing peroxidase: a computational, spectroscopic and site-directed mutagenesis study, Biochem J, 2015, 466, 253–62.

References

| 127

[21] Strittmatter E, Liers C, Ullrich R et al. First crystal structure of a fungal high-redox potential dye-decolorizing peroxidase: substrate interaction sites and long-range electron transfer, J Biol Chem, 2013, 288, 4095–102. [22] Morales M, Mate MJ, Romero A, Martinez MJ, Martinez AT, Ruiz-Duenas FJ. Two oxidation sites for low redox potential substrates: a directed mutagenesis, kinetic, and crystallographic study on Pleurotus eryngii versatile peroxidase, J Biol Chem, 2012, 287, 41053–67. [23] Ogola HJ, Kamiike T, Hashimoto N et al. Molecular characterization of a novel peroxidase from the cyanobacterium Anabaena sp. strain PCC 7120, Appl Environ Microbiol, 2009, 75, 7509–18. [24] Wei K, Luo S-W, Fu Y, Liu L, Guo Q-X. A theoretical study on bond dissociation energies and oxidation potentials of monolignols, Journal of Molecular Structure: THEOCHEM, 2004, 712, 197–205. [25] Jing D. Improving the simultaneous production of laccase and lignin peroxidase from Streptomyces lavendulae by medium optimization, Bioresour Technol, 2010, 101, 7592–7. [26] Santos A, Mendes S, Brissos V, Martins LO. New dye-decolorizing peroxidases from Bacillus subtilis and Pseudomonas putida MET94: towards biotechnological applications, Appl Microbiol Biotechnol, 2014, 98, 2053–65. [27] Tuor U, Wariishi H, Schoemaker HE, Gold MH. Oxidation of phenolic arylglycerol beta-aryl ether lignin model compounds by manganese peroxidase from Phanerochaete chrysosporium: oxidative cleavage of an alpha-carbonyl model compound, Biochemistry, 1992, 31, 4986–95. [28] Roberts JN, Singh R, Grigg JC, Murphy ME, Bugg TD, Eltis LD. Characterization of dyedecolorizing peroxidases from Rhodococcus jostii RHA1, Biochemistry, 2011, 50, 5108–19. [29] Kishi K, Kusters-van Someren M, Mayfield MB, Sun J, Loehr TM, Gold MH. Characterization of manganese(II) binding site mutants of manganese peroxidase, Biochemistry, 1996, 35, 8986–94. [30] Mester T, Field JA. Characterization of a novel manganese peroxidase-lignin peroxidase hybrid isozyme produced by Bjerkandera species strain BOS55 in the absence of manganese, J Biol Chem, 1998, 273, 15412–7. [31] Singh R, Grigg JC, Qin W, Kadla JF, Murphy ME, Eltis LD. Improved manganese-oxidizing activity of DypB, a peroxidase from a lignolytic bacterium, ACS Chem Biol, 2014, 8, 700–6. [32] Kuan IC, Johnson KA, Tien M. Kinetic analysis of manganese peroxidase. The reaction with manganese complexes, J Biol Chem, 1993, 268, 20064–70. [33] Banci L, Camarero S, Martinez AT et al. NMR study of manganese(II) binding by a new versatile peroxidase from the white-rot fungus Pleurotus eryngii, J Biol Inorg Chem, 2003, 8, 751–60. [34] Tien M, Ma D. Oxidation of 4-methoxymandelic acid by lignin peroxidase. Mediation by veratryl alcohol, J Biol Chem, 1997, 8912–7. [35] Letoffe S, Heuck G, Delepelaire P, Lange N, Wandersman C. Bacteria capture iron from heme by keeping tetrapyrrol skeleton intact, Proc Natl Acad Sci U S A, 2009, 106, 11719–24. [36] Sugano Y, Matsushima Y, Tsuchiya K, Aoki H, Hirai M, Shoda M. Degradation pathway of an anthraquinone dye catalyzed by a unique peroxidase DyP from Thanatephorus cucumeris Dec 1, Biodegradation, 2009, 20, 433–40. [37] Li J, Liu C, Li B, Yuan H, Yang J, Zheng B. Identification and molecular characterization of a novel DyP-type peroxidase from Pseudomonas aeruginosa PKE117, Appl Biochem Biotechnol, 2012, 166, 774–85. [38] Zubieta C, Joseph R, Krishna SS et al. Identification and structural characterization of heme binding in a novel dye-decolorizing peroxidase, TyrA, Proteins, 2007, 69, 234–43. [39] Liers C, Bobeth C, Pecyna M, Ullrich R, Hofrichter M. DyP-like peroxidases of the jelly fungus Auricularia auricula-judae oxidize nonphenolic lignin model compounds and high-redox potential dyes, Appl Microbiol Biotechnol, 2010, 85, 1869–79.

128 | 6 Bacterial dye-decolorizing peroxidases

[40] Johjima T, Ohkuma M, Kudo T. Isolation and cDNA cloning of novel hydrogen peroxidedependent phenol oxidase from the basidiomycete Termitomyces albuminosus, Appl Microbiol Biotechnol, 2003, 61, 220–5. [41] Kaur A, Van PT, Busch CR et al. Coordination of frontline defense mechanisms under severe oxidative stress, Mol Syst Biol, 2010, 6, 393. [42] Maurer LM, Yohannes E, Bondurant SS, Radmacher M, Slonczewski JL. pH regulates genes for flagellar motility, catabolism, and oxidative stress in Escherichia coli K-12, J Bacteriol, 2005, 187, 304–19. [43] Liu X, Du Q, Wang Z et al. Crystal structure and biochemical features of EfeB/YcdB from Escherichia coli O157: ASP235 plays divergent roles in different enzyme-catalyzed processes, J Biol Chem, 2011, 286, 14922–31. [44] Dailey HA, Septer AN, Daugherty L et al. The Escherichia coli protein YfeX functions as a porphyrinogen oxidase, not a heme dechelatase, MBio, 2011, 2, e00248–11. [45] Sturm A, Schierhorn A, Lindenstrauss U, Lilie H, Bruser T. YcdB from Escherichia coli reveals a novel class of Tat-dependently translocated hemoproteins, J Biol Chem, 2006, 281, 13972–8. [46] Singh R, Eltis LD. The multihued palette of dye-decolorizing peroxidases, Arch Biochem Biophys, 2015, 574, 56–65. [47] Rahmanpour R, Bugg TD. Assembly in vitro of Rhodococcus jostii RHA1 encapsulin and peroxidase DypB to form a nanocompartment, FEBS J, 2013, 280, 2097–104. [48] Contreras H, Joens MS, McMath LM et al. Characterization of a Mycobacterium tuberculosis nanocompartment and its potential cargo proteins, J Biol Chem, 2014, 289, 18279–89. [49] Sutter M, Boehringer D, Gutmann S et al. Structural basis of enzyme encapsulation into a bacterial nanocompartment, Nat Struct Mol Biol, 2008, 15, 939–47. [50] Dunford HB. Horseradish peroxidase: structure and kinetic properties, in Everse J, Everse KE, Grisham MB, eds. Peroxidases in chemistry and biology, Boca Raton, FL: CRC press, 1991. [51] Chen et al. Characterization of dye-decolorizing peroxidase (DyP) from Thermomonospora curvata reveals unique catalytic properties of A-type DyPs. J Biol Chem, 2015, 290, 23447– 23463. © the American Society for Biochemistry and Molecular Biology. [52] Dunford HB. Heme Peroxidases. New York: John Wiliey, 1999. [53] Hiner AN, Martinez JI, Arnao MB et al. Detection of a tryptophan radical in the reaction of ascorbate peroxidase with hydrogen peroxide, Eur J Biochem, 2001, 268, 3091–8. [54] Sivaraja M, Goodin DB, Smith M, Hoffman BM. Identification by ENDOR of Trp191 as the freeradical site in cytochrome c peroxidase compound ES, Science, 1989, 245, 738–40. [55] Wang X, Peter S, Kinne M, Hofrichter M, Groves JT. Detection and kinetic characterization of a highly reactive heme-thiolate peroxygenase compound I, J Am Chem Soc, 2012, 134, 12897–900. [56] Goodwin DC, Yamazaki I, Aust SD, Grover TA. Determination of rate constants for rapid peroxidase reactions, Anal Biochem, 1995, 231, 333–8. [57] Schonbaum GR, Lo S. Interaction of peroxidases with aromatic peracids and alkyl peroxides. Product analysis, J Biol Chem, 1972, 247, 3353–60. [58] Renganathan V, Gold MH. Spectral characterization of the oxidized states of lignin peroxidase, an extracellular heme enzyme from the white rot basidiomycete Phanerochaete chrysosporium, Biochemistry, 1986, 25, 1626–31. [59] Wariishi H, Akileswaran L, Gold MH. Manganese peroxidase from the basidiomycete Phanerochaete chrysosporium: spectral characterization of the oxidized states and the catalytic cycle, Biochemistry, 1988, 27, 5365–70. [60] Zubieta C, Krishna SS, Kapoor M et al. Crystal structures of two novel dye-decolorizing peroxidases reveal a beta-barrel fold with a conserved heme-binding motif, Proteins-Structure Function and Bioinformatics, 2007, 69, 223–33.

References

|

129

[61] Veitch NC. Horseradish peroxidase: a modern view of a classic enzyme, Phytochemistry, 2004, 65, 249–59. [62] Holm L, Rosenstrom P. Dali server: conservation mapping in 3D, Nucleic Acids Res, 2010, 38, W545–9. [63] Goodin DB, McRee DE. The Asp-His-Fe triad of cytochrome c peroxidase controls the reduction potential, electronic structure, and coupling of the tryptophan free radical to the heme, Biochemistry, 1993, 32, 3313–24. [64] Nonaka D, Wariishi H, Welinder KG, Fujii H. Paramagnetic 13C and 15N NMR analyses of the push and pull effects in cytochrome c peroxidase and Coprinus cinereus peroxidase variants: functional roles of highly conserved amino acids around heme, Biochemistry, 2010, 49, 49–57. [65] Sundaramoorthy M, Terner J, Poulos TL. The crystal structure of chloroperoxidase: a heme peroxidase–cytochrome P450 functional hybrid, Structure, 1995, 3, 1367–77. [66] Singh R, Grigg JC, Armstrong Z, Murphy ME, Eltis LD. Distal heme pocket residues of B-type dye-decolorizing peroxidase: arginine but not aspartate is essential for peroxidase activity, J Biol Chem, 2012, 287, 10623–30. [67] Beitlich T, Kuhnel K, Schulze-Briese C, Shoeman RL, Schlichting I. Cryoradiolytic reduction of crystalline heme proteins: analysis by UV-Vis spectroscopy and X-ray crystallography, J Synchrotron Radiat, 2007, 14, 11–23. [68] Sundaramoorthy M, Gold MH, Poulos TL. Ultrahigh (0.93A) resolution structure of manganese peroxidase from Phanerochaete chrysosporium: implications for the catalytic mechanism, J Inorg Biochem, 2010, 104, 683–90. [69] Geng J, Dornevil K, Davidson VL, Liu A. Tryptophan-mediated charge-resonance stabilization in the bis-Fe(IV) redox state of MauG, Proc Natl Acad Sci U S A, 2013, 110, 9639–44. [70] Bornscheuer UT, Huisman GW, Kazlauskas RJ, Lutz S, Moore JC, Robins K. Engineering the third wave of biocatalysis, Nature, 2012, 485, 185–94. [71] Reetz MT. Laboratory evolution of stereoselective enzymes: a prolific source of catalysts for asymmetric reactions, Angew Chem Int Ed Engl, 2010, 50, 138–74. [72] Reetz MT. Biocatalysis in organic chemistry and biotechnology: past, present, and future, J Am Chem Soc, 2013, 135, 12480–96. [73] Bugg TD, Rahmanpour R. Enzymatic conversion of lignin into renewable chemicals, Curr Opin Chem Biol, 2015, 29, 10–7. [74] Bugg TD, Ahmad M, Hardiman EM, Rahmanpour R. Pathways for degradation of lignin in bacteria and fungi, Nat Prod Rep, 2011, 28, 1883–96. [75] Ornston LN, Stanier RY. The conversion of catechol and protocatechuate to beta-ketoadipate by Pseudomonas putida, J Biol Chem, 1966, 241, 3776–86. [76] Harwood CS, Parales RE. The beta-ketoadipate pathway and the biology of self-identity, Annu Rev Microbiol, 1996, 50, 553–90. [77] Sainsbury PD, Hardiman EM, Ahmad M et al. Breaking down lignin to high-value chemicals: the conversion of lignocellulose to vanillin in a gene deletion mutant of Rhodococcus jostii RHA1, ACS Chem Biol, 2013, 8, 2151–6. [78] Mycroft Z, Gomis M, Mines P, Law P, Bugg TD. Biocatalytic conversion of lignin to aromatic dicarboxylic acids in Rhodococcus jostii RHA1 by re-routing aromatic degradation pathways, Green Chem, 2015. [79] Linger JG, Vardon DR, Guarnieri MT et al. Lignin valorization through integrated biological funneling and chemical catalysis, Proc Natl Acad Sci U S A, 2014, 111, 12013–8. [80] Vardon DR, Franden MA, Johnson CW et al. Adipic acid production from lignin, Energy & Environmental Science, 2015, 8, 617–28. [81] Stolz A. Basic and applied aspects in the microbial degradation of azo dyes, Appl Microbiol Biotechnol, 2001, 56, 69–80.

130 | 6 Bacterial dye-decolorizing peroxidases

[82] Sugano Y, Matsushima Y, Shoda M. Complete decolorization of the anthraquinone dye Reactive blue 5 by the concerted action of two peroxidases from Thanatephorus cucumeris Dec 1, Appl Microbiol Biotechnol, 2006, 73, 862–71. [83] Kandelbauer A, Gubitz GM. Bioremediation for the decolorization of textile dyes – a review, in Lichtfouse E, Schwarzbauer J, Robert D, eds. Environmental chemistry. Berlin: Springer, 2005, pp. 269–88. [84] Mendes S, Farinha A, Ramos CG, Leitao JH, Viegas CA, Martins LO. Synergistic action of azoreductase and laccase leads to maximal decolourization and detoxification of model dyecontaining wastewaters, Bioresour Technol, 2011, 102, 9852–9. [85] Jia J, Wang B, Wu A, Cheng G, Li Z, Dong S. A method to construct a third-generation horseradish peroxidase biosensor: self-assembling gold nanoparticles to three-dimensional sol-gel network, Anal Chem, 2002, 74, 2217–23. [86] Sezer M, Genebra T, Mendes S, Martins L, Todorovic S. A DyP-type peroxidase at a biocompatible interface: structural and mechanistic insights, Soft Matter, 2012, 10314–21. [87] Smulevich G, Spiro TG. Nanosecond transient resonance Raman spectra of the FeII-CO and FeIII-NO photolysis products of horseradish peroxidase, Biochim Biophys Acta, 1985, 830, 80–5. [88] Murgida DH, Hildebrandt P. Electron-transfer processes of cytochrome C at interfaces. New insights by surface-enhanced resonance Raman spectroscopy, Acc Chem Res, 2004, 37, 854–61.

Pei-Ching Chang, Hsi-Yen Hsu, and Guang-Way Jang

7 Biological routes to itaconic and succinic acids 7.1 Introduction There is an ever-increasing interest in bio-based chemicals and materials, due in part to concerns raised by the availability of resources, environmental pollution, and future societal development. Among renewable energies that can be derived from various sources, renewable biomass from photosynthesis is the only sustainable carbon source for most bio-based chemicals and materials. Utilization of agricultural and food processing waste for the production of renewable chemicals results in substantial cost reduction for biorefinery and is also beneficial to the existing agricultural and food industries. The United States Department of Energy (DOE) has identified top value-added building block chemicals that possess various functionalities with rich chemistry and are suitable for multiple transformations into a wide range of intermediates and materials. US biorefinery-related initiatives focus on renewable energy and bulk material production to reduce petroleum dependency, a strategic good. Potential benefits of white biotechnology advancements for the US include national security, reduced GHG emissions, an increase in the number of carbon-fixing plants, and rural development. On the other hand, the initial focuses of the EU are the manufacture of novel, high-margin products to revive global competitiveness of its chemical industry and initiate, as urged by a few active national NGOs, a reduction of the EU’s carbon footprint. Succinic acid and itaconic acid have been identified by the US Department of Energy in 2004 as top value-added chemicals. Both chemicals are commercially available, with annual global production capacity thereof at 30 000 to 50 000 tons each. Succinic acid is a naturally occurring chemical that has been identified in bovine rumen, a CO2 -rich environment. At present, most industrial applications of succinic acid are produced via chemical routes. On the other hand, itaconic acid is currently manufactured using biological methods. Succinic acid and itaconic acid are two platform chemicals with very similar molecular structures, distinguished only by the additional unsaturated double bond on the 2-position for that of itaconic acid. After a decade of development, many bio-based succinic acid plants are in operation, with still more planned or under construction, while production of itaconic acid is frequently stalled due to limited applications thereof.

7.1.1 Succinic acid Succinic acid (IUPAC systematic name: butanedioic acid; traditionally known as amber acid) is a versatile platform chemical with applications in both high-value

132 | 7 Biological routes to itaconic and succinic acids

niche personal care and food additives markets and in the large volume production of polyester, polyurethanes, plasticizers, and coatings. At present, succinic acid is predominantly produced from butane through catalytic hydrogenation of petroleumbased maleic anhydride. According to a researchandmarkets projection, the succinic acid market will reach $ 486.7 million by 2019 with a CAGR of 22.6 % by volume between 2014 and 2019 [1]. The global succinic acid production capacity is between 30 000 and 50 000 tons at a compound annual growth rate of 18.7 % from 2011 to 2016. Based on succinic acid applications, the global succinic acid market can be segmented into the following sectors and their respective market share percentages: industrial applications, 57.1 %; pharmaceuticals, 15.91 %; food & beverages, 13.07 %; others, 13.92 % (PRNewswire, June 16, 2015). Succinic acid is one of the US Department of Energy’s 12 top value-added chemicals from biomass. The estimated potential global market for succinic acid is expected to reach US $ 7–10 billion per year assuming that the price of succinic acid production will be substantially reduced in the future. Recognizing the potential of bio-based succinic acid, companies interested in manufacturing this C4 platform chemical include BioAmber Inc., Myriant Corporation, Reverdia (a DSM-Roquette joint venture), Succinity (a BASF-Purac joint venture), Agro-Industrie Recherches et Développements (ARD), Mitsui & Co., China National BlueStar Co. and PTT Public Company Limited. Bio-succinic acid plants around the globe are summarized in Tab. 7.1 [2]. Succinic acid is a platform chemical most widely used in food ingredients, as precursors to active pharmaceutical ingredients and pharmaceutical additives and has the potential in industrial applications to replace maleic anhydride and serve as a precursor for the production of 1,4-butanediol (BDO), tertahydrofuran (THF), N-methyl pyrrolidone (NMP) and γ-bytyrolactone (GBL). 1,4-Butanediol can be further carbonylated to synthesize adipic acid for the manufacture of nylon 6,6, lubricants, foams and food products. BDO is also raw material for the preparation of polybutylene terephthalate (PBT), a high performance plastic. Succinates can be used as additives to animal feed and as precursors for protein synthesis. Diethyl succinate can be used for paint stripping and ethylene diaminedisuccinate is a replacement for EDTA. Other applications for succinic acid and derivatives thereof include catalysis of food seasoning preparation, cut flower preservation, soil chelation, corrosion inhibition and modification of polyester resin to improve dyeing ability. The largest existing succinic acid market involves its use as a surfactant, detergent extender and foaming agent [3]. At present, most succinic acid for industrial use is produced by petrochemical process and only natural derivatives thereof utilized in the food market are produced by fermentation. Through biological route, succinic acid is produced as an intermediate of the tricarboxylic acid cycle or as an end product of anaerobic metabolism. The leading developers of the field, BioAmber and Reverdia utilize genetically modified yeast whereas Myriant applied E. coli strains for the biosynthesis of succinic acid. Biosuccinic acid is generally considered safe. A list of maximum-level use for succinic acid in desserts, soups and broths, as well as that for starch sodium octenyl succinate

7.1 Introduction

| 133

Tab. 7.1: Global bio-succinic acid plants (planned and in production) [2]. Company

Capacity (tons/y)

Plant location

Operational date

Microorganism

Downstream

Basfia succiniciproducens

Mg-based process

Yeast

Electrodialysis

E. coli

Ammonia precipitation

yeast

direct crystallization

BASF-Purac JV

50 000

TBA*

TBA*

BASF-Purac JV

25 000

Barcelona, Spain

2013

BioAmber-ARD

3 000

Pomacle, France

Full capacity by Q2 2012

BioAmber-Mitsui JV

65 000

TBA* (US or Brazil)

TBA*

BioAmber-Mitsui JV

30 000

Sarnia, Ontario, Canada

2015-08-06

BioAmber-Mitsui JV

65 000

Thailand

2014

Myriant

13 600

Lake Providence, Louisiana

Q1 2013

Nanjing, China

TBA*

Lake Providence, Louisiana

Q1 2014

Infraleuna site, Germany

H1 2012

Cassano Spinola, Italy

H2 2012

Myriant-China National BlueStar

110 000

Myriant

Myriant-Uhde (owner and operator) Reverdia (DSM-Roquette)

77 110

500

10 000

in weaning food, can be found in directive 95/2/EC [4]. Bio-succinate has also been used as a flavoring enhancer for low-sodium food and emulsifiers in infant formulas and follow-on formulas. In pharmaceutical applications, succinate is used as an anticarcinogenic and an insulinotropic agent [4, 5]. There is a wide range of applications for succinic acid and derivatives thereof [3, 6].

7.1.2 Itaconic acid Itaconic acid, 2-methylidenebutanedioic acid, is an unsaturated di-carbonic acid and has been used for the production of acrylic plastics, acrylate latexes, alkyl paints, industrial adhesives, superabsorbents, detergents, and antiscaling agents. At present, synthetic latex is the largest application accounting for 55 % of itaconic acid in the

134 | 7 Biological routes to itaconic and succinic acids

global market. The unsaturated vinylidene functionality of itaconic acid renders the bio-based chemical with a reactive site for generating unique molecular structures or producing a polymer with improved UV-resistant characteristics. Itaconic acid is emerging as a prospective bio-based replacement of maleic anhydride for the preparation of unsaturated polyester resins (UPRs) due to similarities between the two chemicals. Recent developments in itaconic acid applications have been in biomedicine, including those in ophthalmology, dentistry and drug delivery [7]. Chemical conversion of citric acid to itaconic acid by controlled pyrolysis/distillation processes has being described [8]. The low price difference between citric and itaconic acids limits the economic efficiency of the chemical routes. Currently, most manufacturers are producing itaconic acid with Aspergillus terreus using pretreated molasses under phosphatelimited conditions. Global Industry Analysts, Inc. forecasts the global itaconic acid market to reach US $ 398.3 million by 2017. In particular, the Asia-Pacific region is poised to become the fastest growing itaconic acid market at 9 % CAGR, while the United States represents the largest itaconic acid market. The key players in the global market are highly concentrated in China.

7.2 Synthesis of succinic acid and derivatives Succinic acid is a C4 -dicarboxylic acid with important applications in a wide range of industries including those of food, chemicals, plastics, and pharmaceuticals. As a CO2-fixing process, the biological route to succinic acid is considered environmentally benign. It is suggested that succinate fermentations have the potential to reach the production volumes of citric acid or even ethanol [3]. Thus, production of biosuccinic acid can generate significant new markets for agricultural carbohydrates. Recent economical assessments comparing biological routes to chemical processes suggest a cost competitiveness of bio-based succinic acid over that conventionally derived from petrochemicals [9]. The estimated total production cost is € 2554/MT of succinic acid produced by the hydrogenation of maleic anhydride, followed by hydration of the resulting intermediate succinic anhydride to synthesis succinic acid. On the other hand, estimated costs for the production of succinic acid by the fermentation methods are between € 85 and € 1233/MT of succinic acid depending on the type of biomass feedstock and technique applied. Fermentative succinic acid production surpassed that of petrochemical production in 2013. The success of producing bio-based succinic acid on an industrial scale depends on its potential as an alternative to maleic anhydride and as an intermediate for the synthesis of 1,4-butandiol (BDO) and polybutylene succinate (PBS). Forecasted market demand for succinic acid in 2020 range from 500 000–700 000 tons to 2 million tons [6, 10, 11]. Energy efficiency calculations also suggest that fermentative succinic acid production is preferable to that of petrochemical production. In case of a poor sugar conversion of 70 % efficiency for a second generation lignocellulosic biomass, the trend is reversed. Key challenges facing

7.2 Synthesis of succinic acid and derivatives |

135

the production of bio-based succinic acid are material efficiency (product mass/input mass) and E factor (waste mass/product mass). Material efficiency and E factor for petrochemical succinic acid production processes using maleic anhydride are 76 % and 0.3, respectively. A large quantity of water is required for the fermentation process. The two factors can be improved by recovery of water and valorization of byproduct ammonium sulfate (Myriant process) or coproduction of ethanol (Reverdia process).

7.2.1 Chemical synthesis of succinic acid Succinic acid for current industrial applications is predominantly produced by the hydrogenation of petroleum-based maleic acid or maleic anhydride. The raw material, maleic anhydride, is obtained by catalytic oxidation of C4 hydrocarbons. Petrochemical production of succinic acid is achieved by catalytic hydrogenation of maleic anhydride to prepare succinic anhydride followed by hydration. The resulting solution is then concentrated to enable crystallization of succinic acid at low temperature. The product is recovered by centrifugation and drying. For the synthesis of 1,4-BDO from maleic anhydride, succinic acid is produced as an in situ intermediate. Separation and drying are thus irrelevant. Alternative chemical routes to succinic acid include paraffin oxidation, electrolytic reduction of maleic anhydride in acidic medium, catalytic addition of acetylene and acrylic acid [5, 12]. Electrochemical synthesis has the advantage of high product purity, which is especially useful in food and pharmaceutical applications.

7.2.2 Biological routes to succinic acid Current biological routes to produce succinic acid on a commercial scale employ E. coli and yeast strains. As with many biomass-derived chemicals, bio-succinic acid is still disadvantaged in terms of cost competitiveness when compared to its petroleumbased counterparts; feedstock price and the processing cost are key considerations. A broad range of carbon and nitrogen sources have been studied as potential feedstocks. Molasses, sugar mixtures, and glycerol were evaluated as candidates for carbon sources, while yeast extracts and wheat processing byproducts were used as nitrogen sources. The downstream isolation and purification process are critical to the production cost. It was suggested that the price of succinic acid needs to fall below US $ 0.45 per kilogram to open up commodity markets thereof [3]. According to Wilke’s model, it will require 100 % (w/w) yield, a productivity of 3 g/l/h, and titers up to 250 g/l to achieve the target production cost [13, 14]. A very limited number of succinic acid biosynthesis processes achieve a productivity of greater than 3 g/l/h [13]. Such productivity strongly governs plant capacity and thus affects both variable costs and fixed costs, while yield on feedstock and feedstock price have direct im-

136 | 7 Biological routes to itaconic and succinic acids

pact on variable costs. Considering future projection quantities and cost, continuous integrated production is likely to outperform batch process for the biosynthesis of succinic acid. Fermentative production of succinic acid from renewable resources is not only advantageous in terms of cost effectiveness, but also results in CO2 fixation. However, the biological route requires considerable space and amount of water, as well as long fermentation time and a complicated product isolation process [12]. Fermentative succinate-producing microorganisms can be generally classified into natural (bacteria, fungi) and engineered (bacteria, yeast) species [5, 13]. Natural bacteria species show high tolerance to osmotic pressure caused by high succinate concentration, but their cultivation requires expensive nutrients. Fungal strains, on the other hand, show lower productivity than that of their bacterial counterparts. Notable microorganisms engineered for efficient succinate production are E. coli, Corynebacterium glutamicum, and S. cerevisiae. Strain engineering strategies to enhance succinate productivity include inactivation of branch pathways, overexpression of genes directly involved in branch pathways, redirection of metabolic flux, and the introduction of reducing power. Biosynthesis of succinate can proceed through anaerobic and aerobic conditions. Anaerobic fermentation is preferred over aerobic fermentation because of its low capital investment and operational cost. Three formation pathways for succinate are the glyoxylate path and the oxidative and the reductive branches of the triarboxylic acid (TCA) cycle [15]. The glyoxylate and the oxidative pathways are active under aerobic conditions whereas the reductive branch of the TCA cycle proceeds under anaerobic conditions. NADH generation in the glyoxylate route alone results in an imbalance of electrons. Activation of the glyoxylate pathway in conjunction with anaerobic fermentation improves succinate yield [16]. The succinate yield and byproduct formation in anaerobic culture are strongly governed by the availability of NADH. It is critical to utilize genetic engineering tools to manipulate metabolic pathways to achieve a suitable intracellular redox balance. During the oxidative branch of the TCA cycle, the conversion of succinate to fumarate should be blocked. The theoretical yield is 1.71 moles of succinate produced per mole of glucose in the presence of CO2 . Such a theoretical yield increases to 2 moles of succinate produced per mole of glucose with electron donor supplements, such as hydrogen [17]. The nature of the fermentation pathway and CO2 /H2 ratio are critical factors in succinate production yield. Succinate is one key intermediate of the TCA cycle. Thus, it is possible to synthesize succinic acid by evaluating metabolic pathways leading to other TCA cycle intermediates, such as lactate, acetate, and ethanol, and genetically engineering microorganisms accordingly. Microorganisms for the production of bio-based succinic acid include Actinobacillus succinogenes, Anaerobiospirillum succiniciproducens, Mannheimia succiniciproducens, C. glutamicum, Bacteroides fragilis, E. coli, and Yarrowia lipolytica [6, 15, 18, 19]. The physiology of succinate-producing microorganisms varies significantly from one species to the next [3]. For example, both

7.2 Synthesis of succinic acid and derivatives | 137

A. succiniciproducens and A. succinogenes follow the phosphoenolpyruvate (PEP) carbocykinase pathway exclusively to produce succinate. A. succiniciproducens is an obligate anaerobe, while A. succinogenes is a facultative anaerobe, able to switch between aerobic and anaerobic metabolic pathways. On the other hand, E. coli, a facultative anaerobe, proceeds through multiple pathways to produce succinate. A. succinogenes, unlike A. succiniciproducens and E. coli, has high tolerance to succinate salts. The PEP carboxykinase pathway is regulated by CO2 concentration to a selected product distribution. Under high CO2 levels (100 mol CO2 /100 mol glucose), succinate is the major product, with traces of lactic acid or ethanol. At a low CO2 level, A. succiniciproducens produces lactic acid as a major product, whereas A. succinogenes generates more ethanol than succinic and lactic acids. Carboxylation of pyruvate, the most important central metabolite, is the controlling step of succinic acid biosynthesis. Typical biosynthesis of succinic acid utilizes glucose or glycerol as a carbon source while assimilating CO2. Succinic acid can be produced in anaerobic and aerobic conditions [15]. Succinate yield through a fermentative pathway is limited by the supply of NADH. A combined anaerobic and aerobic strategy is often applied for the biosynthesis of succinic acid. Carbon sources, carbon dioxide, hydrogen and culture pH are critical factors for economic production of succinic acid. In addition to external CO2 gas, carbonates are considered as a source of CO2. Carbon dioxide and carbonates dissolved in water generate HCO−3 and CO−3 . The equilibrium between CO2, HCO−3 , and CO−3 is governed by pH in fermentation broth. In a biosynthesis system, hydrogen is a potential electron donor to improve succinate yield by decreasing cellular redox potential and improving NADPH recycles. In addition to phosphoenolpyruvate, oxaloacetate, and malate, pyruvate was identified as a fourth node governing A. succinogenes fermentation pathways to succinate and alternative products [20]. At high CO2 and H2 concentrations, A. succinogenes produces more succinate and less formate, acetate, and ethanol. Studies suggest high NaHCO3 concentrations minimized the tendency to push the flux from the C4 pathway to C3 pathway. Most fermentation processes are carried out in batch reactors. High titers and yields of succinate, however, were often obtained in fed-batch or continuous systems. Succinate titers of 133 g/l were reported by using engineered C. glutamicum in a fermentation medium consisting of glucose, bicarbonate and formate under anaerobic condition (Fig. 7.1). At present, biosynthetic processes have limited potential for industrial scale application due to the use of formate which complicates downstream purification and increases production cost [6]. Okino et al. achieved a very high succinic acid titer of 146 g/l with 1.40 mol/mol yield under oxygen deprivation with intermittent addition of sodium bicarbonate and glucose [21]. Succinate titers of up to 110 g/l at 83–87 wt.% yield were achieved with A. succinogenes by maintaining the pH with magnesium [12, 22]. The recombinant E. coli AFP184 (Fig. 7.2) demonstrated productivity of 3 g/l/h by applying dual-phase fermentations in high sugar concentrations whereas E. coli produced succinic acid with titers of up to 99.2 g/l at productivities

138 | 7 Biological routes to itaconic and succinic acids

ATP

Glucose PEP

Pyruvate Glucose 6-phosphate ATP

2 NADH CO2

PEP

Lactate

Oxaloacetate

Pyruvate

NADH NADH

Formate Malate

Acetyl-CoA

Fumarate Acetate

Ethanol

Succinate

Fig. 7.1: Simplified metabolic pathway of wild-type C. glutamicum for succinate production. PEP, phosphoenolpyruvate [21].

Glucose PEP ATP Pyruvate Glucose 6-phosphate 2 NADH CO2

ATP

PEP

Lactate

Oxaloacetate NADH

Pyruvate

Formate Malate

Fumarate

NADH

Acetyl-CoA

Acetate

Ethanol

Succinate

Fig. 7.2: Simplified anaerobic production of succinate in E. coli AFP184. The blocked reactions are shown by crosses [25].

7.2 Synthesis of succinic acid and derivatives | 139

of up to 1.3 g/l/h [12]. A very high productivity of 10.4 g/l/h at a titer of 83 g/l and a 1.35 mol/mol yield were obtained in a continuous system with a membrane for cell recycling and an on-line electrodialysis system to eliminate the end-product inhibition [15, 23]. Productivity reaches 14.8 g/l/h at a titer of 42 g/l when using an integrated membrane-bioreactor-electrodialysis system under a suitable culture condition [23]. The integrated fermentation system produces a highly concentrated cell-free succinate solution that can be recovered by simple water evaporation/acidification. The production yield affects the variable costs of raw materials and utilities while productivity and titer govern fixed costs and total production investment? Low production rate and low product concentrations result in high energy input and cost. Downstream recovery and purification represent both technological and economical challenges. Product isolation processes will need to be coupled with upstream fermentation to render industrial scale-up manufacturing viable. Strategies include genetically engineering strains to reduce acid byproducts, reactive extraction to improve succinate selectivity, in situ product removal, and separation of acetic acid from succinic acid using an emulsion liquid membrane (ELM) [15, 24]. Thus, to improve the competitiveness of succinic acid biosynthesis, it is essential to address the importance of succinatetolerant microorganisms, low-cost biomass feedstocks and downstream product recovery. The succinate fermentation efficiencies of a selected number of bacteria strains are summarized in Tab. 7.2. An ideal microorganism candidate for industrial production must be able to utilize a wide range of carbon sources, such as fructose, glucose, sucrose, maltose, lactose and xylose, for the synthesis of succinic acid. This capability provides the opportunity Tab. 7.2: Summary of succinate fermentation efficiency using different bacteria strains. Strain

Corynebacterium glutamicum ΔldhA-pCRA717

Titer of succinic acid (g/l)

Fermentation conditions

146

Escherichia coli 99.2 AFP111/pTrc99Apyc; ΔpflAB::CmR, ldhA::KmR, ptsG– , pyc+ Actinobacillus 94–106 succinogenes mutants Anaerobiospirillum 50.3 succiniciproducens

Productivity

Ref.

(g/l/h)

Yield of succinic acid (g/g)

Under oxygen deprivation with repeated intermittent addition of glucose and sodium bicarbonate Dual phase (aerobic to anaerobic)

3.2

0.92

[21]

1.3

1.1

[10]

Batch

2.0–2.8

0.83–0.87

[3]

Batch; anaerobic

2.1

0.90

[10]

140 | 7 Biological routes to itaconic and succinic acids

to employ agricultural and food processing wastes as feedstock to reduce production cost. However, E. coli almost exclusively uses glucose for succinic acid production. Wang et al. engineered an E. coli strain SBS550MG bearing both the pHL413 plasmid and the pUR400 plasmid for utilizing sucrose and mixed sugars to produce succinate [26]. Evaluation of different substrates indicates that the nature of sugar carbon sources affects the fermentation pathways and thus the yield and productivity of succinate. The E. coli strain SBS550MG pHL413 produced the highest succinate yield of 1.86 mol/mol with a productivity of 0.82 g/l/h when provided with fructose as a carbon source. The strain consumed glucose preferentially and grew better in a mixture of glucose and fructose than in fructose alone. The succinate yield was 1.76 mol/mol hexose with a productivity of 1.21 g/l/h in a glucose/fructose mixture of 1/1 ratio. E. coli are not able to consume sucrose to produce succinate due to the lack of an invertase. E. coli strains able to metabolize sucrose were achieved via expression of the pUR400 plasmid containing scrK, Y, A, B, and R genes that can convert sucrose to β-Dfructose and α-D-glucose 6-phosphate. However, a low productivity of 0.48 g/l/h was obtained using SBS550MG pHL413 pUR400 due to slow sucrose hydrolysis during the early stage of the fermentation process. The fermentation strategy was then changed to utilize sucrose during the aerobic phase before switching to a mixture of fructose, glucose, and sucrose during the anaerobic phase. The processes achieved a yield of 1.67 mol/mol hexose with a productivity of 0.85 g/l/h. The strain consumed sugars in the following priority: glucose, sucrose, and then fructose. Fermentation in a sucrose hydrolysis solution achieved a similar result, with a succinate yield of 1.67 mol/mol hexose and a productivity of 0.86 g/l/h. Studies on the effect of the dissolved oxygen (DO) level during the aerobic phase suggest a high agitation rate is conducive for increased cell growth rate and succinate productivity, while low agitation is better for increased succinate production yield. Formation of a microaerobic condition before switching to a completely anaerobic condition (due to rapid depletion of the DO level following slower agitation of the growth medium) may have contributed to differences in cell metabolism and led to improved succinate yield. Li et al. utilized corn stover hydrolysate as a carbon source and spent yeast hydrolysate as a nitrogen source to produce 56.4 g/l succinic acid with A. succinogenes [27]. They achieved 0.73 g/g yield and over 50 % production cost reduction. Earlier studies indicated that the hydrolysis of corn stover generated a 2 : 1 (w/w) ratio of glucose to xylose. A mixed sugar feedstock resulted in delayed succinic acid production when compared with that achieved with glucose only. In general, consumption of xylose starts only when glucose is depleted. A wide range of carbon sources including corn starch, corn steep liquor, whey, cane molasses, glycerol, and lignocelluloses, have been evaluated for the fermentative production of succinic acid. Metabolic engineering approaches to facilitate simultaneous digestion of sugars are beneficial for the industrial production of bio-succinate. The delay in succinic acid production resulting from mixed sugar feedstocks may have also been caused by the presence of inhibitors, such as furfural, HMF, and acetic acid, in corn stover hydrolysate. E. coli is an ideal candidate for future commercial bio-

7.2 Synthesis of succinic acid and derivatives |

141

succinate production due to in-depth knowledge on the genetic manipulation thereof, a high possible yield of 1.72 mol/mol glucose, and the general acceptance of its usage in industrial process [16]. Applying solid-state fermentation (SSF) with two fungi strains and using wheat milling byproducts, Dorado et al. were able to produce enough carbon and nitrogen nutrients for the production of succinic acid [28]. Agricultural and food wastes are more easily used in solid-state fermentation (SSF) than in submerged fermentation. Other low cost raw materials used for succinic acid fermentation include corncobs, corn straw and stalk, rapeseed meal, cassava starch, spirit-based distiller grains, and cane molasses [18, 29]. Typical strains employed for the production of succinic acid include isolated microorganisms, recombinant microorganisms and or improved strain (screened mutations). Biomass derived media and fermentation strategies affect the yield and productivity of bio-based succinic acid production. Genetic engineering and fermentation strategies provide a range of economic approaches for flexible and integrated production. Succinic acid was co-produced with other value-added products, such as malic acid [30], isoamyl acetate [31] and polyhydroxybutyrate [32, 33]. For fresh water conservation, seawater was successfully used for the fermentation of wheatderived substrates to produce succinic acid [34]. Separation of succinic acid from fermentation broth accounts for a major fraction of succinic acid production cost. Typical isolation methods include precipitation, reactive extraction, and direct crystallization while integrated membrane filtration-electrodialysis is the most promising approach with improved titer and productivity. Utilization of agro-waste and integration of processes results in significant cost reduction. It is possible to produce succinic acid by fermentation for about US $ 0.55–1.10 per kg according to McKinlay et al. [17]. Producing succinic acid with biological routes is expected to save up to 40 % energy consumption and result in lower carbon emissions when compared to production by typical chemical processes [29]. Life cycle assessment of bio-succinic acid prepared by low pH yeast-based fermentation technology and direct crystallization downstream processing indicate a reduction of 90 % non-renewable energy use and reduced greenhouse gas (GHG) emissions by a factor of two when compared to succinic acid prepared with petro-based maleic anhydride [35]. Downstream recovery and purification account for considerable costs when producing succinic acid. The separation process includes removal of cells and proteins, concentration of succinate, conversion of succinic salt to acid, and purification of succinic acid [3, 5]. Downstream processing cost can as account for up to 60–70 % of the total succinic acid production cost [12]. At first, microbial cells are separated from the fermentation broth by centrifugation or filtration followed by ultrafiltration to remove proteins, polysaccharides, and other oligomers. Several approaches to isolate succinate from fermentation broth were evaluated [3, 17]. Succinic acid can be obtained by isolating succinate from fermentation broth by electrodialysis followed by a conversion process using a bipolar, water-splitting membrane stack. The resulting succinic acid can then be purified by ion exchange chromatographies to achieve

142 | 7 Biological routes to itaconic and succinic acids

80 % recovery in dry weight. The remaining 20 % of the recovered material consists of acetic acid. Datta et al. described in their patent for a precipitate succinate hydroxide process to maintain the pH of the fermentation broth in the range of 5.8 to 6.4 by addition of calcium oxide or calcium hydroxide [36]. The resulting aqueous slurry of calcium succinate was then transformed to succinic acid by introducing sulfuric acid. The resulting product was then purified with ion exchange resin to obtain succinic acid containing less than 1 % nitrogenous impurities. The disadvantage of this process is that it requires handling large amounts of slurry and calcium sulfate waste. Alternatively, the pH of the fermentation broth can also be maintained by adding NH4 OH and later reacting the resulting diammonium succinate with ammonium bisulfate to form ammonium sulfate and succinic acid. Addition of sulfic acid to this mixture of ammonium sulfate and succinic acid resulted in the crystallization of succinic acid. The succinic acid was then recovered in 90 % dry weight by dissolution in methanol to precipitate contaminating sulfates. Succinic acid was also purified by reactive extraction with tri-n-octylamine. There is still a need for considerable efforts to lower the recovery cost of succinic acid to achieve economically viable industrial scale production thereof.

7.2.3 Catalytic conversion of succinic acid Typical chemical transformations of succinic acid include esterifications, amidations and hydrogenations. Esterifications of succinic acid were used to prepare diester intermediates for polymerization. High esterification yields of greater than 75 % were achieved for low purity succinic acid recovered from fermentation broths by means of an approach combining both conductive heating and microwave irradiation [37]. This in situ esterification in fermentation broth was also utilized as a method for downstream isolation of succinic acid. Catalysts for the in situ transformation process must be tolerant to water and salts, as well as resistant to organic contaminates. Hydrogenation of succinic acid produces a range of value-added chemicals, including tetrahydrofuran (THF) (used as a solvent), γ-bytyrolactone (GBL) (used as a solvent), as an intermediate in the manufacture of pyrrolidones, as an ingredient in the production of pesticides and herbicides, and 1,4-butanediol (BDO) for polymer synthesis. Direct downstream catalytic transformation of succinic acid into valuable derivatives in filtered aqueous fermentation broth eliminates the need for isolation and thus reduces production cost [38]. Petrochemicals are mostly in a low oxidation state and therefore require oxidative derivatization, whereas renewable chemicals are often in a high oxidation state and need to be reduced. Hydrogenation of maleic acid results in the formation of succinic acid. Thus, hydrogenation of maleic acid in the aqueous phase can be easily adapted to produce succinic acid in a reduction process. Group VIII metals are the most active catalysts for the hydrogenation of succinic acid to produce γbutyrolactone (GBL), tetrahydrofuran (THF), and 1,4-butanediol (BDO). Combinations

7.2 Synthesis of succinic acid and derivatives |

143

of Group VIII metals, or their combinations with other metals, such as rhenium and tin, are often deposited on carbon, TiO2 and ZrO2 substrates to serve as catalysts. In addition to the nature of metals and properties of supports, dispersion and repartition of the metals on and in their catalyst supports are also critical for the catalytic efficiency of the hydrogenation process. A high degree of dispersion in bimetallic catalysts prevents the formation of unwanted microstructures that lead to catalyst aging and loss of activity. On the other hand, reaction parameters have a strong impact on product distribution and yield. Reductive amination of succinic acid using amine, ammonium, or ammonia produces pyrrolidones and derivatives. 2-Pyrrolidone is an intermediate in the preparation of nylon-4, pharmaceuticals, medicines and agrochemicals. N-methyl-2-pyrrolidone (NMP) is an important solvent in the polymer and electronic industries. The reaction selectivity appears to be sensitive to the reactant ratios and an excess of alcohol is required for the production of NMP. Direct aqueous synthesis of pyrrolidones from succinic acid without producing GBL was achieved [38]. Catalytic conversion of succinic acid into valuable derivatives is illustrated in Fig. 7.3. O O O

H2

O

O MAn (Petrochem.)

OR

RO

O H2

O Succinic ester OH HO 1,4-Butanediol (BDO)

O Succinic anhydride

O

O

O

Amine

O

NH

O HO

OH

Γ-Butyrolactone (GBL)

H2

O Succinic acid (Fermentation)

O

HO

Tetrahydrofuran (THF)

n PTMG

O NH

H2+ CH3NH2

NCH3 N-methyl Pyrrolidone

2-Pyrrolidone

O

H2+ NH3

or

2-Pyrrolidone

O NCH3

N-methyl

Pyrrolidone

Fig. 7.3: Catalytic transformation of succinic acid into valuable derivatives.

Nylon4

H

144 | 7 Biological routes to itaconic and succinic acids

The hydrogenation of succinates is similar to the Davy–Mackee process for the commercial production of 1,4-butanediol from maleate. Copper-based catalysts were studied for the hydrogenation of succinates to produce BDO [39, 40]. Alternatively, the hydrogenation of succinic anhydride over Cu–Zn, Cu–Zn–Zr, Cu–Zn–Cr–Zr, Cu–Mn, Re–Cu–Zn was performed using vaper-phase fixed-bed rectors [41–46]. Ruthenium complexes were utilized as homogeneous catalysts for the hydrogenation of succinic anhydride to GBL under mild conditions [47–50]. Other homogeneous catalysts include Ru acetylacetonate, trioctylphosphine and HBF4 [51–53]. Although Ru complexes provide high product selectivity, drawbacks of using homogeneous catalysts include difficulty of separation from the reaction medium and the presence of unfavorable halogen ligands. Aqueous phase hydrogenation of succinic acid is environmentally benign and enjoys reduced production costs. However, acidic media poses a challenge for metal catalysts. Ly et al. evaluated Pd–Re bimetal catalysts with various Re content for aqueous phase hydrogenation [54]. It was observed that the Re position and oxidation states (Re3+ , Re0 ) on the reduced bimetallic samples were related to their preparation procedures [55]. The presence of a small amount of Re in a Pd/C catalyst enhances production of γ-butyrolactone, whereas further increase of Re loading resulted in a high THF yield [56]. Additional studies also indicate hydrogenation efficiencies can be improved by using palladium-rhenium catalysts supported by carbon materials [57, 58]. The modified Ru catalysts also provide high conversions of succinic acid in the aqueous solution to produce THF and 1,4-butanediol over Ru–Mo, Ru–Sn, Ru–Re, and Ru–Re–Sn trimetall catalyst [59–62]. Succinic acid can be transformed into 1,4-butanediol with a yield of over 70 % in the presence of a Pd-5FeOx /C catalyst under the relatively mild conditions of 200 °C and 5 MPa H2 [63]. The supporting substrate plays an important role in governing catalyst particle size and thus hydrogenation performance. A Pd/SiO2 –NH2 catalyst prepared by cocondensation method exhibits high catalytic activity in converting succinic acid to γ-butyrolactone with 100 % conversion and 94 % selectivity using 1,4-dioxane as a solvent at 240 °C and 60 bar for 4 h [64]. Kim et al. evaluated the catalytical activity of Pd-WO3 /Al2 O3 for hydrogenation reactions in a mixture selected from dioxane, ethanol, and H2 O at 100–300 °C [69]. Studies of supported ruthenium and rehenium catalysts treated with acid for the hydrogenation of succinic acid indicate that the yields of THF increase as catalyst particle size decreases [65–68]. It was identified that mesoporous carbon and Alumina xerogel provide improved hydrogenation efficiency due to the fine dispersion of catalyst on these supports [69, 70]. A very small amount of Pt (100 ppm) in gold catalysts facilitates H2 dissociation and achieves 97 % selectivity to GBL at 97 % conversion [71]. The composition of Ru-Co bimetallic catalyst governs the reaction pathway and therefore the distribution of products [72].

7.3 Synthesis of itaconic acid and derivatives | 145

7.3 Synthesis of itaconic acid and derivatives Itaconic acid currently occupies only a niche market with somewhat tenuous market expansion potential with suppliers limited to an extremely narrow region. Itaconic acid, a structurally unique C5 bio-based chemical, has an α, β-conjugated methylene group and two differentiated carboxylate functionalities from which a diverse range of chemicals and materials for commercial applications may be derived [73–77]. Examples include polyester, thermoplastics, elastomers, coatings, adhesives, artificial glass, superabsorbent polymers, bioactive compounds with agricultural applications, pharmaceuticals, and biomedical applications [78]. At present, the itaconic acid market is small, with current annual global production a little more than 80 000 tons [79]. Further expansion of the itaconic acid market requires a reduction in its production costs and an increase in the flexibility in scale-up itaconic acid production operations. The currently employed itaconic acid chemical synthesis route is impractical because of its high cost and low yield [75]. At present, Aspergillus terreus is a strain being used for commercial production of itaconic acid [80]. As A. terreus is a fungus, there are limited genetic engineering tools for pathway manipulation and process improvement for its fermentation. Previous studies have shown that CAD is an essential enzyme for itaconate biosynthesis [80–86]. However, amino acid sequencing for the CAD enzyme was not completed until 2008 [87]. Instead of utilizing A. terreus, improvement in itaconate production efficiency via fermentation is more feasible by means of engineering an E. coli synthetic pathway. This is because E. coli is a well-characterized microorganism with a set of readily available tools for genetic manipulation and its physiological regulation is well-studied.

7.3.1 Biological routes to itaconic acid Although some Aspergillus species and other microorganisms were identified as candidates to produce itaconic acid, A. terreus remains the dominant host for industrial itaconic acid production [88]. Biosynthesis of itaconic acid is an aerobic process requiring 1.5 mol of O2 for every mole of bio-based diacid generated (Fig. 7.4) [8]. It is, therefore, a challenge to balance between the competing concerns of sufficient oxygen to the growth medium and limiting cell damage of filamentous microorganisms, such as A. terreus, resulting from the hydromechanical stress of aeration. The influence was reduced to some degree by using an air lift bioreactor and suitable control of medium pH. High itaconic acid yields were obtained by fermentation of glucose. For commercial production, product yield per substrate is critical for profitability. Theoretically, 1 g of glucose can be converted to 0.72 g itaconic acid. Yahiro et al. achieved yields of up to 0.57 g itaconic acid per gram of glucose, but observed a reduction in productivity and yield at titers above 60 g/l [90]. Subsequent optimization strategies to gain

146 | 7 Biological routes to itaconic and succinic acids

Itaconate

Glucose Extracellular Cytosol

IA exporter Cis-aconitate

Fructose 6-phosphate

CAD

Itaconate

MTT Pyruvate

Pyruvate

Cis-aconitate

Isocitrate

Citrate Oxaloacetate

Malate

Mitochondrion

Acetyl-CoA Oxaloacetate

α-ketoglutarate TCA cycle Succinyl-CoA

Malate Fumarate

Succinate

Fig. 7.4: Biosynthesis of itaconic acid in A. terreus [89]. MTT, mitochondrial tricarboxylate transporter.

cost competitiveness are enhancing productivity and titer as well as using low cost substrates. Studies indicate some limitation on the chemical mutation of A. terreus to achieve high titer. It has been proposed that the transfer into A. terreus of certain genes in more robust microbial producers (such as A. niger which is known to produce high yields and titers of citric acid) may lead to higher titers. Productivity of industrial batch fermentation using molasses as a substrate is in the range of 1 g/l/h [8]. To be competitive with petroleum-based chemicals, productivity of a typical biosynthesis process needs to reach at least 2.5 g/l/h. Continuous fermentations may help achieve high fermentation productivity. Various sugars, such as glucose, sucrose, xylose, saccharose, lactose, starch, molasses, and hydrolysate derived from lignocellulosic materials, can be carbon sources for the biosynthesis of itaconic acid using A. terreus. Utilization of wood waste as a feedstock for production of bio-based acids proves less efficient, due to the presence of inhibitory compounds; making matters worse, xylose leads to lower yields than those of other sugars. A process to separate inhibitors in the biomass hydrolysate might be inevitable. A. terreus can also utilize pure glycerol for the production of itaconic acid at yields of up to 55.9 wt.% after 233 hours [8]. In addition to A. terreus and A. niger, non-filamentous microorganisms, such as Pseudozyma anatarctica and U. maydis, various yeasts, such as Y. lipolytica, and bacteria, such as E. coli, were evaluated for the production of itaconic acid with some degree of success. Methods for downstream recovery and purification of itaconic acid include crystalliza-

7.3 Synthesis of itaconic acid and derivatives | 147

tion, reactive extraction using organophosphorous compounds or quaternary amine, and electrodialysis. There is a critical step to separate itaconic acid and residual glucose which is known to interfere with the crystallization. Bred strains of A. terreus can secrete a significant concentration of itaconate (> 80 g/l) in a media. However, this is still far from the titers in excess of 200 g/l achieved in the citric acid industrial process and the estimated maximum theoretical titer of 240 g/l [91]. Although the itaconic acid production pathway is not clear, CAD was found to be the key enzyme of the pathway. A high transcriptional level of the cad gene is crucial for the efficiency of itaconic acid biosynthesis. Due to poor stability of the CAD protein, the amino acid sequencing thereof was not completed until a substantial amount of the enzyme was purified in 2008. In addition to screening existing strains, high titers can be achieved by employing targeted genetic engineering. On the other hand, media compositions and engineering approaches related to oxygen distribution and bioreactor design also contribute to itaconic acid production efficiency. Introduction of copper ions and steady aeration were found to positively influence itaconic acid production. Various reactors including bubble column reactors, tubular reactors, packed bubble column reactors, and Air-Lift reactors have been evaluated for biosynthesis. Starch was utilized as a carbon source for fermentation to reduce production cost. Hydrolysis of starch with glucoamylase (5000 AUN/ml) or nitric acid at pH 2 led to itaconic acid yields of 0.36 and 0.35 g/g starch, respectively [7]. Relatively high titers were also achieved with agricultural residuals, such as rejected bananas (28.5 g/l), apples (31.0 g/l), Jatropha seed cakes (24.45 g/l), and sago starch (48.2 g/l). Chemical routes for itaconic acid synthesis are impractical, while fermentative itaconic acid production using A. terreus is economically challenging [80]. Utilization of itaconic acid, therefore, is limited by its high production cost and limited number of suppliers [78]. To lower itaconate production cost, fermentation was enhanced by genetic manipulation and physiological regulation of E. coli. Okamoto et al. achieved 4.34 g/l itaconic acid production by icd inactivation and acnB overexpression in cad-expressing E. coli [86]. They also developed a recombinant E. coli expressing α-amylase on its cell surface to saccharify starch near the cell surface and utilize the resulting sugar for the biosynthesis of itaconic acid [92]. A novel synthetic pathway in E. coli to produce itaconate is shown in Fig. 7.5. Itaconate is produced by a submerged-cultured fermentation with a strain of E. coli in a medium containing a carbohydrate source such as glycerol, glucose, or molasses. Nutrients contain amino acids, phosphates and metal salts, as well as sulfuric acid, hydrochloric acid or sodium hydroxide can be added as pH adjustment agent. A medium containing 6 wt.% of glycerol is incubated at 30–35° and pH of 6.0–7.0 for 48 hours. Itaconate was produced at 1.5 g/l/h and a 99.5 wt.% pure product was produced in a continuous purification section, which includes double-effect evaporators, two-stage crystallization, activated carbon treatment, and rotary dryer operation units. Successful production of itaconate from the engineered E. coli opens up the possibility of using non-native, easily manipulated organisms for itaconate production [93]. Eco-

148 | 7 Biological routes to itaconic and succinic acids Itaconate cad Acn Cis-aconitate Acn Isocitrate Citrate

Glucose Pyruvate PEP

Ppc

Acetyl-CoA

gltA Oxaloacetate

Icd α-ketoglutarate TCA cycle Succinyl-CoA

Malate Fumarate

Succinate

Fig. 7.5: Schematic representation of itaconate production in engineered E. coli (ppc, phosphenolpyruvate carboxylase; gltA, citrate synthase; acn, aconitase; cad, cis-aconitic acid decarboxylase; PEP, phosphoenolpyruvate; OAA, oxaloacetate) [93].

nomic analyses of a fermentation process using E. coli yielded a production cost of $ 2.0/kg for an itaconate plant with annual capacity of 20 000 MT. The calculation of such a production cost was based on experimental results of 80 g/l titer and 1.5 g/l/h productivity, as well as on assumptions of a 95 % recovery rate, crude glycerol cost of $ 0.2/kg, a nutrient cost of $ 0.5/kg and a 10-year plant life.

7.3.2 Catalytic conversion of itaconic acid The catalytic conversions of itaconic acid to 2-methyl-1,4-butanediol, 3-methyltetrahydrofuran, and 2-methyl γ-butyrolactone as well as 3-methyl γ-butyrolactone should be similar to those associated with the transformation of succinic acid to related derivatives [94–100]. The hydrogenation of itaconic acid to 2-methylsuccinic acid, a chemical with pharmaceutical applications, was performed with Ru-Starbon, Ru/TiO2 or Ru-complex as a catalyst [100, 101].The enantioselective hydrogenation of itaconic acid to (R)- or (S)-methylsuccinic acid using palladium and rhodium catalysts is also described. Decarboxylation of itaconic acid to bio-based methacrylic acid (MAA) was achieved with transition-metal catalysts in aqueous phase [102, 103]. Catalytic conversion of succinic acid into valuable derivatives is illustrated in Fig. 7.6.

7.4 Applications of succinic acid and derivatives Industrial uses for succinic acid include its applications as a surfactant, an ion chelator for corrosion inhibition, a flavoring agent and pH regulator in food processing, and as an ingredient for the production of pharmaceutical materials, such as antibiotics, amino acids, and vitamins [12]. It is also a platform chemical for the synthesis of adipic

7.4 Applications of succinic acid and derivatives |

CH3

O

OR

RO

H2

O

O Itaconic ester

O H3C

3-methyl GBL

CH3

4-methyl GBL

H3C

OH

O

HO 2-methyl BDO

3-methyl THF

O

O OH

HO

–CO2

Methyl methacrylic acid

OH

O Itaconic acid

O

O

H3C

149

O

NH3

NH2

H2N

Itaconic diamide

O

H3C

O

O

NCH3

NCH3

Reductive amination

3-methyl NMP

CH3 H2N

H3C

4-methyl NMP

H3C

NH2 2-methyl –1, 4-butanediamine

NCH3 3-methyl pyrrolidine

Fig. 7.6: Catalytic transformation of succinic acid into valuable derivatives.

acid, 1,4-butanediol, tetrahydrofuran, N-methyl pyrrolidinone, 2-pyrrolidinone, γ-butyrolactone, succinic acid esters, and succinate salts. Polyesters, polyamides, and polyester amides can be synthesized by using succinic acid and its diamine and diol derivatives. Polybutylene succinate (PBS) prepared by the polycondensation of succinic acid and 1,4-butanediol has superior processability, biodegradability, and balanced mechanical properties. The physical properties of PBS resemble those of a conventional plastic, polyethylene (PE); thus, the bioplastic has great potential to be used for the fabrication of grocery bags, packaging films, and mulch films. Polybutylene succinate is typically prepared by transesterification polymerization consisting of transesterification and polycondensation steps using tetran-butyl-titanate or tetraisopropyl titanate as a catalyst. Lipase was applied for the synthesis of PBS under mild conditions without using metal salts [12]. In general, esterification takes place at a temperature range between 150–200 °C under a low vacuum followed by polycondensation at temperature between 220–240 °C under a high vacuum [12]. It is essential to have proper water removal and temperature control during esterification and a sufficiently high vacuum during polycondensation; it is also

150 | 7 Biological routes to itaconic and succinic acids

necessary to use a catalyst with high reactivity and resistance to hydrolysis in order to obtain PBS of high molecular weight. Mochizuki et al. obtained a PBS with molecular weight (Mn) of 59 000 by using tetra-n-butoxy titanate as a catalyst and polyphosphoric acid as a thermal stabilizer [12, 104]. Alternatively, polymerization can be performed in solution at relatively low temperatures to avoid oxidation of PBS. A series of catalysts including SnCl2 , Sn(Oct)2 , Ti(OiPr)4 , Ti(OBu)4 , Zn(Ac)2 , and p-TS (p-toluenesulfonate) were evaluated for the synthesis of PBS by solution polymerization, with SnCl2 identified as the most effective in receiving high molecular polymers [12]. PBS of high molecular weight can also be achieved by using chain extenders after condensation polymerization. Diisocyanate and anhydride are often selected for the chain extension of hydroxyl-terminated PBS while oxazoline and epoxy are appropriate for enhancing the molecular weight of carboxyl-terminated PBS. Showa Denko’s Bionolle is an example of high molecular weight PBS prepared by coupling condensed prepolymers using hexamethylene diisocyanate as a chain extender. Branching increases the melt strength and thus enhances the workability of a polymer in processes involving elongated flow, such as fiber spinning, film blowing, vacuum forming, and foaming [12]. Copolymerization and branching strategies provide alternatives to tailordesigned physical properties and biodegradability of PBS. PBS and copolymers are semicrystalline polyesters and their crystalline structures vary with copolymer composition. Foreseeing the potential of bioplastics, PBS manufacturers are becoming more numerous and currently include Hexing Chemical, Xinfu Pharmaceutical, BASF, Eastman, Mitsubishi Chemicals, and SK Chemicals [12]. Bio-based thermoplastic polyester elastomers (TPEE) were obtained by catalystfree polycondensation of renewable chemicals, including glycerol, azelaic acid and succinic acid [105]. Glycerol is a byproduct of biodiesel manufacture while azelaic acid is commercially available by the oxidative cleavage of oleic acid from palm oil. The resulting bio-based TPEEs demonstrated superior thermal stability. Development of the biocompatible TPEEs is targeting orthopedic and ophthalmic applications, as well as those in reconstructive surgery and drug delivery. Thermogravimetric studies indicated the primary degradation occurs at temperatures above 400 °C. The glass transition temperature of the resulting TPEE increases with increasing succinic acid content from about –23 °C for TPEEs containing a 1:1 ratio of glycerol and azelaic acid to about –8 °C for TPEEs containing equal amounts of azelaic and succinic acids.

7.5 Applications of itaconic acid and derivatives Water soluble polyitaconic acid may be used as superabsorbents, antiscaling agents in water treatments, co-builders in detergents, and dispersants for minerals in coatings. Additional applications can be identified by copolymerization with other monomers, such as hydrogels, latex and elastomers. Copolymers of itaconic acid with acrylic and methacrylic acid as well as their esters can be used for the preparation of coatings and

7.5 Applications of itaconic acid and derivatives |

151

adhesives. Direct polymerization of itaconic acid with a redox initiator in aqueous or dioxan media leads to products of complicated structures [106]. It is suggested that formation of anhydride and decarboxylation occurred during radical polymerization and drying because only about half of the acid groups were detected by potentiometric titration. FTIR studies suggest decarboxylation occurs when the polymer is dried at 120 °C. Alternatively, polymerization of itaconic anhydride followed by hydrolysis of the resulting poly(itaconic anhydride) led to a poly (itaconic acid) consisting of two carboxylic acids in a single monomeric unit [107]. A deep eutectic solvent (DES) consisting of a 1:1 ratio of itaconic acid and choline chloride (2-hydroxyethyltrimethyl ammonium chloride, vitamin B4) was selected as the polymerization medium for the preparation of poly(itaconic acid) hydrogels using ammonium persulfate as a redox initiator and N,N’-methylenebisacrylamide as a crosslinking agent [108]. These studies concluded that polymerization kinetic of itaconic acid was enhanced and crosslinking density was higher in DES than that in water. The author suggests that association of choline chloride with carboxylic acids via hydrogen bonding and catalyzation of persulfate decomposition by ammonium salts may contribute to the acceleration of polymerization. The presence of a methylene group alone with dicarboxylic functionality distinguishes itaconic acid from other biomass derived diacids. Thermoplastic polyester elastomers (TPEEs) consist of alternating crystalline hard segments and amorphous soft segments [109]. The crystalline fractions, as physical crosslinks resembling chemical crosslinking regions in vulcanized rubber, provide strength to elastomers. The amorphous segment governs the softness and resilience of elastic materials. Thermoplastic polyester elastomers are often used to replace traditional natural rubber and thermosetting polymers due to their superior physical and chemical properties, such as tensile strength, elastic recovery, impact resistance, creep resistance, cold resistance, flexural fatigue resistance, oil resistance, and solvent resistance. Applications of TPEEs include those in engineering plastics, automotives, cushioning, packaging, sports equipment and medical equipment [110]. DuPont Hytrel and DSM Anitel ECO are TPEEs prepared from bio-based chemicals. Bio-based TPEEs can also be achieved by using itaconate derivatives for the preparation of TPEEs soft segments (Fig. 7.7). Dimethyl 2-methyl succinate (DM2MS) and 2-methyl butane diol (2m-BDO) were obtained by the hydrogenation of itaconic acid [111]. Condensation of the two monomers followed by polycondensation of the resulting IA-based polyol with PBT segments was used to prepare an IA-based TPEE. As shown in Tab. 7.3, IA-based TPEEs demonstrate compatible mechanical properties with commercial TPEEs and the elongation of biobased elastomers increases along with an increase of polyol content, whereas tensile strength and hardness (Shore-D) values decrease. The degree of crystallinity of hard segments is critical to the performance of the resulting IA-based TPEE. Dicarboxylic acid sodium salt was used as a nucleating agent to enhance crystallinity of IA-based TPEE. The elastomer is amorphous at 200 °C. As observed with a polarized optical microscope, crystallization occurs as temperature drops below its crystallization temper-

152 | 7 Biological routes to itaconic and succinic acids O

Fermentation

Glucose/ glycerol

HO

Catalyst

HO

O Itaconate Catalyst

CH3 O

O

OH

O O

IA-Polyol CH3

–H2O

O

O

O

OH

CH3 OR

O Dialkyl 2-methyl succinate CH3

Block Polymerization

HO

O PBT segment

OH

2-methyl BDO

O HO

H2

RO

n

O HO

OR

RO

O Itaconic acid CH3

O

ROH

OH

IA -TPEE (ITRI) O

O

O

O

O n

O

O

O O

O

PBT segment

CH3

CH3 O

m

IA-Polyol Fig. 7.7: Schematic representing synthetic routes of IA-based TPEE.

Tab. 7.3: Physical properties of IA-based TPEE as a function of polyol loading. IA-polyol (wt.%)

Intrinsic viscosity (IV)

Tensile strength (kgf/cm2)

Elongation (%)

Shore-D

50 45 40 35

1.02 1.01 0.99 1.03

101 110 115 135

963 805 630 560

28 35 44 50

Tab. 7.4: Structural changes observed as a function of nucleation agent concentration. Hard/soft segment

Nucleated agent (in situ)

Tensile (kgf/cm2)

Elongation (%)

Crystallinity ΔH (J/g)

Shore-D

6:4 6:4 6:4

— NU-1 (2000 ppm) NU-1 (5000 ppm)

115 136 165

630 595 740

16.5 16.9 19.0

44 46 46

7.6 Conclusions

|

153

ature, Tc . Crystal growth rates of the elastomers with nucleating agent concentrations of 2000 and 5000 ppm at 100 °C are 0.30 and 0.67 µm/minute, respectively. Mechanical performance of IA-based TPEE improved as the crystallinity increased (Tab. 7.4). An additional merit of these new IA-based TPEEs is their exceptional UV resistance. This is because an IA-based polyol consists of an ester group and tertiary substituted carbon. The two adjacent units in the elastomer backbone form a stable radical thus resisting polymer degradation upon UV irradiation. The radical created by UV radiation is stabilized and quenched in the sequential IA-polyol units eliminating cleavage along the main chain [112, 113]. IA-based TPEEs maintain mechanical properties even after 200 hours of UV light irradiation, whereas commercial TPEEs became brittle and fractured after comparable UV exposure. Bio-based polyamides were synthesized by utilizing organic salts of itaconic acid and diamines in the presence of a sodium dihydrogen phosphate (NaH2 PO4 ) catalyst to prevent blanching and crosslinking of the itaconic acid moiety [114]. Formation of a rigid N-substituted pyrrolidone ring in the polymer main chain renders biopolyamides with improved mechanical strength and environmental corrosion characteristic via a ring opening reaction. The biomass-derived polyamides have comparable thermal degradation temperature to those of polyhexamethylene adipamide (PA66) and polycaprolactam (PA6), whereas their glass transition temperatures of 80–97 °C are much higher than those of the two conventional polyamides. Although the rigid pyrrolidone ring causes steric hindrance for interchain hydrogen bonding, biopolyamides demonstrate a superior mechanical strength (σ) of 90–165 MPa and Young’s modulus (E) of 430–2800 MPa, as compared to conventional polyamides. As-prepared biopolyamides are not soluble in water and adsorb less than 4 % water. However, the biopolyamides are soluble in alkaline solution due to pyrrolidone ring-opening hydrolysis and corroded in soil with a pH between 7.5 and 7.9.

7.6 Conclusions The development of efficient and cost-effective biorefineries using agricultural waste and inedible plant parts is essential for achieving a sustainable economic, ecological, and social system. Both succinic acid and itaconic acid consist of a C4 back bond with di-carboxyl acid and can be produced via biological routes with high efficiency. In addition to fermentation, research efforts are also focused on downstream processing, as product recovery costs account for up to 60–70 % of the total production cost. Itaconic acid possesses an unsaturated vinyldine side chain that provides reactivity and UV-resistant characteristics for products or intermediates containing the moiety. Industrial synthesis of itaconic acid is currently based on a biological route using A. terreus. Applications of itaconic acid, at present, focus on specialty markets, such as those for latexes, TPEE, polyamides, paints, industrial adhesive, superabsorbents, detergents, and antiscaling agents. Potential pharmaceutical application has also been

154 | 7 Biological routes to itaconic and succinic acids

explored. However, large-scale applications of itaconic acid have yet to be identified. The feasibility of utilizing itaconic acid for the production of methyl methacrylate (MMA) could be a critical factor governing future market demand. Succinic acid was predominantly produced from petrochemicals for applications in high-value niche markets of personal care and food additives as well as in large volume products of polyester, polyurethanes, plasticizers and coatings. To achieve high production efficiency, a combined anaerobic and aerobic strategy is often applied for the biosynthesis of succinic acid. A very high succinic acid titer of 146 g/l with 1.40 mol/mol yield was achieved under oxygen deprivation with intermittent additions of sodium bicarbonate. An on-line electrodialysis system was utilized to limit end-product inhibition and obtain a succinic acid productivity of 10.4 g/l/h at titer of 83 g/l and a 1.35 mol/mol yield. Production capacity of bio-succinic acid exceeded succinic acid production via a petroleum route in 2013 due to its competitive manufacturing cost. Although current market demand is limited to about 50 000 tons, there is potential to utilize the platform chemical for the production of BDO, for which the annual global market exceeds three million tons. Acknowledgment: We would like to acknowledge the Ministry of Economic Affairs of Taiwan, Republic of China for financial support. Brian Jang edited and reviewed, and Vivian Fan compiled the text.

References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17] [18]

RESEARCHANDMARKETS. “Succinic Acid Market by Source, Application – Global Trends & Forecast to 2019”, December 2014. Jansen ML, Gulik WMV. Current Opinion in Biotechnology, 2014, 30, 190–197. Zeikus JG, Jain MK, Elankovan P. Appl Microbiol Biotechnol, 1999, 51, 545–552. Chimirri F, Bosco F, Ceccarelli R, Venturello A, Geobaldo F. Ital J Food Sci, 2010, 22, 119–125. Cao Y, Zhang R, Sun C, Cheng T, Liu Y, Xian M. BioMed Research International, Volume 2013, Article ID 723412 Leszczewicz M, Walczak P. Biotechnol Food Sci, 2014, 78, 25–43. Hajuan H, Yusoff WMW. Curr Res J Biol Sci, 2015, 7, 37–42. Klement T, Büchs J. Bioresource Tech, 2013, 135, 422–431. Pinazo JM, Domin M. E, Parvulescu V, Petru F. Catalysis Today, 2015, 239, 17–24. McKinlay JB, Vieille C, Zeikus JG. Appl Microbiol Biotechnol, 2007, 76, 727–740. Roland Berger Strategy Consultants, Bioplastics Market Study, August 2012. Xu J, Guo BH. Microbiol Monographs, 2010, 14, 347–388. Beauprez JJ, De Mey M, Soetaert WK. Process Biochemistry, 2010, 45, 1103–1114. Wilke D. Appl Microbiol Biotechnol, 1999, 52, 135–145. Cheng KK, Zhao XB, Zeng J, Zhang JA. Biofuels, Bioprod Bioref, 2012, 6, 302–318. Cheng KK, Wang GY, Zeng J, Zhang JA. BioMed Research International, Volume 2013, Article ID 538790. McKinlay J, Vieille C, Zei J. Appl Microbiol Biotech, 2007, 76, 727–740. Shui Z, Qin H, Wu B, Tan F, Wang J, He M. Chin J Appl Environ Biol, 2015, 21, 10–21.

References

[19] [20] [21] [22] [23] [24] [25] [26] [27] [28] [29] [30] [31] [32] [33] [34] [35] [36] [37] [38] [39] [40] [41] [42] [43] [44] [45] [46] [47] [48] [49] [50] [51] [52] [53] [54] [55]

|

155

Isar J, Agrawal L, Saran S, Kaushik R, Saxena RK. Anaerobe, 2007, 13, 50–56. McKinlay JB, Vieille C. Metab Eng, 2008, 10, 55–68. Okino S, Noburyu R, Suda M, Jojima T, Inui M, Yukawa H. Appl Microbiol Biotechnol, 2008, 81, 459–464. Guetter MV, Rumler D, Jain MK. Int J Syst Bacteriol, 1999, 49, 207–216. Meynial-Salles I, Dorotyn S, Soucaille P. Biotechnol Bioeng, 2008, 99, 129–135. Lee SC, Kim HCJ. Membrane Sci, 2011, 367, 190–196. Andersson C, Hodge D, Berglund KA, Rova A. Biotechnol Prog, 2007, 23, 381–388. Wang J, Zhu J, Bennett GN, San KY. Metabolic Eng, 2011, 11, 328–335. Li J, Zheng XY, Fang XJ, Liu SW, Chen KQ, Jiang M, Wei P, Ouyang PK. Bioresource Tech, 2011, 102, 6147–6152. Dorado MP, Lin SKC, Koutinas A, Du C, Wang R, Webb C. J Biotechnol, 2009, 143, 51–59. Lin CKS, Luque R, Clark JH, Webb C, Du C. Biofuels, Bioprod Bioref, 2012, 6, 88–104. Jantama K, Haupt MJ, Savoronos S, Zhang X, Moore JC, Shanmugam KT, Ingram L. O Biotech Bioeng, 2008, 99, 1140–1153. Dittrich CR, Bennett GN, San KY. Biotechnol Prog, 2009, 25, 1304–1309. Kang Z, Gao C, Wang Q, Liu H, Qi Q. Bioresour Technol, 2010, 101, 7675–7678. Kang Z, Du L, Kang J, Wang Y, Wang Q, Liang Q, Qi Q. Bioresour Technol, 2011, 102, 6600–6604. Lin CKS, Luque R, Clark JH, Webb C, Du C. Energy Environ Sci, 2011, 4, 1471–1479. Cok B, Tsiropoulos I, Roes AL, Patel MK. Biofuels, Bioprod Bioref, 2014, 8, 16–29. Datta R, Glassner DA, Vick Roy JR. Fermentation and Purification Process for Succinic Acid 1992, US Patent 5,168,055. Luque R, Lin CKS, Du C, Macquarrie DJ, Koutinas A, Wang R, Webb C, Clark JH. Green Chem, 2009, 11, 193–200. Delhomme C, Weuster-Botz D, Kühn FE. Green Chem, 2009, 11, 13–26. Thomas DJ, Stammbach MR, Cant NW, Wainwright MS, Trimm DL. Ind Eng Chem Res, 1990, 29, 204–208.. Turek T, Trimm DL, Black DS, Cant NW. Appl Catal A, 1994, 116, 137. Castiglioni GL, Fumagalli C. WO Patent 9938856, 1999. Tong L, Wang H, Feng W, Gao G, Li X, Deng J, Zhang X. US Patent 5637735, 1997. Lancia R, Vaccari A, Fumagalli C, Armbruster E. WO Patent 9522539, 1995. Taylor PD, De T, Waldo B, Donald W. US Patent 5122495, 1992. Suzuki S, Inagaki H, Ueno H. JP Patent 02233632, 1990. Suzuki S, Inagaki H, Ueno H. EP Patent 373946, 1990. Lyons JE. J Chem Soc, Chem Commun, 1975, 412. Bianchi M, Menchi G, Francalanci F, Piacenti F, Matteoli U,Frediani P, Botteghi C. J Organomet Chem, 1980, 188, 109. Ikariya T, Osakada K, Ishii Y, Osawa S, Saburi M, Yoshikawa S. Bull Chem Soc Jpn, 1985, 157, 897. Inagaki H, Nighimura S, Yoshinori H, Wada K. Science and technology in catalysis, Kodanska Ltd.: Tokyo, 1994, 327. Utsunomiya M, Mizoguchi M, Iwasaka H. WO Patent 2006041038, 2006. Sugiyama H, Takahashi K, Usaka H. EP Patent 676239, 1995. Fuchigami T, Wakasa N, Ka T, Koga K, Myake T. JP Patent 07017960, 1995. Ly BK, Tapin B, Aouine M, Delichere P, Epron F, Pinel C, Especel C, Besson M. ChemCatChem, 2015, 7, 2161–2178. Tapin B, Epron F, Especel C, Ly BK, Pinel C, Besson M. Catalysis Today, 2014, 235, 127–133.

156 | 7 Biological routes to itaconic and succinic acids

[56] [57] [58] [59] [60] [61] [62] [63] [64] [65] [66] [67] [68] [69] [70] [71] [72] [73] [74] [75] [76] [77] [78] [79] [80] [81] [82] [83] [84] [85] [86] [87] [88] [89] [90] [91]

Shao Z, Li C, Di X, Xiao Z, Liang C. Industrial & Engineering Chemistry Research, 2014, 53, 9638–9645. Liang C, Shao Z, Li C, Xiao Z. CN Patent 103113325, 2013. Minh DP, Besson M, Pinel C, Fuertes P, Petitjean C. Topics in Catalysis, 2010, 53, 1270–1273. Campos D. US Patent 20040122242, 2004. Tooley PA, Black JR. US Patent 5985789, 1999. Schwartz JT. US Patent 5478952, 1995. Bockrath RE, Campos D, Schwartz JT, Stimek RT. US Patent 6008384, 1999. Liu X, Wang X, Xu G, Liu Q, Mu X, Liu H. Journal of Materials Chemistry A: Materials for Energy and Sustainability, 2015, 3, 23560–23569. You C, Zhang C, Chen L, Qi Z. Applied Organometallic Chemistry, 2015, 29, 653–660. Hong UG, Kim JK, Lee J, Lee JK, Song JH, Yi J, Song IK. Journal of Industrial and Engineering Chemistry, 2014, 20, 3834–3840 Hong UG, Kim JK, Lee J, Lee JK, Song JH, Yi J, Song IK. Applied Catalysis, A: General, 2014, 469, 466–471. Hong UG, Park HW, Lee JW, Hwang S, Kwak J, Yi J, Song IK. Journal of Nanoscience and Nanotechnology, 2013, 13, 7448–7453. Hong UG, Park HW, Lee J, Hwang S, Yi J, Song IK. Applied Catalysis, A: General, 2012, 415–416, 141–148 Hong UG, Park HW, Lee J, Hwang S, Song IK. Journal of Industrial and Engineering Chemistry, 2012, 18, 462–468. Hong UG, Hwang S, Seo JG, Yi J, Song IK. Catalysis Letters, 2010, 138, 28–33. Budroni G, Corma A. Journal of Catalysis, 2008, 257, 403–408. Deshpande RM, Buwa VV, Rode CV, Chaudhari RV, Mills PL. Catalysis Communications, 2002, 3, 269–274. Werpy T, Petersen G. Top value added chemicals from biomass, in Laboratory TPNNLatNRE, editor. Richland, WA Department of Energy, 2004. Baup S. Ann Chim Phys, 1837, 19, 29. Corma A, Iborra S, Velty A. Chemical Reviews, 2007, 107, 2411–502. Willke T, Vorlop KDB. Appl Microbiol and Biotechnol, 2001, 56, 289–295. Dwiarti L, Yamane K, Yamatani H, Kahar P, Okabe M. J Bioscience and Bioengineering, 2002, 94, 29–33. Determination of market potential for selected platform chemicals, itaconic acid, succinic acid, 2,5-furandicarboxylic acid. Bratislava, Slovakia: WEASTRA. Okabe M, Lies D, Kanamasa S, Park EY. Appl Microbiol Biotechnol, 2009, 84, 597–606. Winskill N. J Gen Microbiol, 1983, 129, 2877–2883. Bentley R, Thiessen CP. J Biological Chemistry, 1957, 226, 673–687. Bentley R, Thiessen CP. J Biological Chemistry, 1957, 226, 689–701. Bentley R, Thiessen CP. J Biological Chemistry, 1957, 226, 703–720. Jaklitsch WM, Kubicek CP, Scrutton MC. J General Microbiology, 1991, 137, 533–539. Bonnarme P, Gillet B, Sepulchre AM. J Bacteriology, 1995, 177, 3573–8. Okamoto S, Chin T, Hiratsuka K, Aso Y, Tanaka Y, Takahashi T, Ohara H. J Gen Appl Microbiol, 2014, 60, 191–197. Kanamasa S, Dwiarti L, Okabe M, Park EY. Appl Microbiol and Biotechnol, 2008, 80, 223–229. Steiger MG, Blumhoff ML, Mattanovich D, Sauer M. Frontiers in Microbiol, 2013, 4, 1–5. Klement T, Büchs J, Bioresource Technology, 2013, 135, 422–431. Yahiro K, Takahama T, Park YS, Okabe M. J Fement Bioeng, 1995, 79, 506–508. Li A, Van Luijk N, Ter Beek M, Caspers M, Punt P, Van der Werf M. Fungal Genet Biol, 2011, 48, 602–611.

References

[92] [93] [94] [95] [96] [97] [98] [99] [100] [101] [102] [103] [104] [105] [106] [107] [108] [109] [110] [111]

[112] [113] [114]

| 157

Okamoto S, Chin T, Nagata K, Takahashi T, Ohara H, Aso Y. J Biosci Bioeng, 2015, 119, 548–553. Liao JC, Chang PC. US Patent 8143036, 2010. Fischer R, Pinkos R, Wulff-Doering J. DE Patent 19720657, 1997. Weyer HJ, Fischer R, Merger F, Frank J, Henkelmann J, Siegel H, Ruehl T. DE Patent 4231782, 1992. Weyer HJ, Fischer R, Merger F, Frank J, Henkelmann J, Siegel H, Ruehl T. US Patent 5536854, 1995. Rao Velliyur NM. US Patent 4782167, 1988 Fischer R, Pinkos R, Wulff-Doring J. US Patent 6204417, 1998. Mou X, Li S, Wang X, Yao S, Peng G, Jiang Y, Guo X, Zhou J. CN patent 104923218, 2015. Besson M, Gallezot P, Pinel C. Chem Rev, 2014, 114, 1827–1870 Huang Q, Yu W, Lu R, Lu F, Gao J, Miao H, Xu J. RSC Adv, 2015, 5, 97256–97263. Nôtre JL, Dijk SCMW, Haveren JV, Scott EL, Sanders JPM. ChemSusChem, 2014, 7, 2712–2720. Carlsson M, Habenicht C, Kam LC, Antal MJJ, Bian N, Cunningham RJ, Jones M. J Ind Eng Chem Res, 1994, 33, 1989–1996 Mochizuki H, Hirami M. Polym Adv Technol, 1997, 8, 203–209. Baharu MN, Kadhum AAH, Al-Amiery AA, Mohamad AB. Green Chemistry Letters Reviews, 2015, 8, 31–38. Stawski D, Polowinski S. Polimery, 2005, 50, 118–122. Polowinski S. Polimery, 2006, 51, 270–275. Bednarz S, Fluder M, Galica M, Bogdal D, Maciejaszek I. J Appl Polym Sci 2014, DOI: 10.1002/APP.40608. Rodriguez E, Katime I. Macromolecular Materials and Engineering, 2003, 288, 607–612. Mayumi J, Nakagawa A, Matsuhisa K, Takahashi H, Iijima M. Polymer J, 2008, 40, 1–9. Besler A, Harward, A, Marquard, W. Reaction networks – A rapid screening method, in Jezowski, J, Thullie, T. editors. 19th European Symposium on Computer Aided Process Engineering, 2009, p. 243–8. Pan JQ, Zhang J. Polym Degrad Stabil, 1992, 36, 65–72. Tabankia MH, Philippart JL, Gardette JL. Polym Degrad Stabil, 1985, 12, 349–362. Ali MA, Tateyama S, Oka Y, Kaneko D, Okajima MK. Macromolecules, 2013, 46, 3719–3725.

Lei Chen, Xingxun Liu, and Ka-Hing Wong

8 Novel nanoparticle materials for drug/food delivery-polysaccharides 8.1 Introduction As condensation polymers, polysaccharides are generally termed as glycans, in which more than ten monosaccharide units are mutually joined together by O-glycosidic bonds [1]. The general formula of polysaccharides can be represented as (C6 H10 O5 )n where 40 ≤ n ≤ 3000 [2, 3]. Polysaccharides are often quite heterogeneous, with structures ranging from linear to highly branched. Polysaccharides are the most abundant resources in nature that are commonly applied in daily life, including cellulose, pectin, chitosan, starch, etc. Depending on the type of monosaccharide building units, polysaccharides can be divided into homopolysaccharide (also called homoglycan, containing the same type of monosaccharide) and heteropolysaccharide (or heteroglycan, composed of more than one type of monosaccharide) based on the definition given in the International Union of Pure and Applied Chemistry (IUPAC). Both homopolysaccharides and heteropolysaccharides may possess homo-linkages or hetero-linkages with respect to configuration and/or linkage position. These macromolecules can also be divided into storage polysaccharides (such as starch and glycogen) and structural polysaccharides (such as cellulose and chitin) based on their biological functions [4]. Starch is made up of a mixture of amylose and amylopectin. The amylose content in starch depends on the biological source, accounting about 20–30 % in most starch [5]. Amylose is linear chain glucan with α-1,4 linkage, whereas amylopectin consists of several side chains on O-6 site of the glucose in α-1,4 backbone with α-1,6-bonds occurring every 24 to 30 glucose units. Starch is used as a storage polysaccharide in plants, while glycogen serves as the form of energy storage in animals and fungi [6]. The structure of glycogen is similar to amylopectin but is more extensively branched and compact than starch [7]. As a hyperbranched biopolymer, glycogen consists of linear glucose chains with side chains branching off every ten glucoses or so. Overall, most of the storage polysaccharides are hyperbranched molecules. As the most common structural polysaccharide, cellulose is the structural component of the primary cell wall of plants and considered as the most abundant natural resource. This type of polysaccharide consist of a linear chain ranging from one hundred to over ten thousand β-1,4 linked D-glucose units [8]. The other common structural polysaccharide is chitin, which is a long-chain polymer of N-acetylglucosamine [9]. Chitin is the main component of the cell walls of fungi, the exoskeletons of arthropods and insects, the radulas of mollusks, and the beaks and internal shells of cephalopods [10]. Chitin plays the role of cellulose in structure, but serves as keratin in function. As

160 | 8 Novel nanoparticle materials for drug/food delivery-polysaccharides

a naturally occurring polymer, chitin has also been proven its usefulness in medical and industrial applications. Some studies have described several health benefits of polysaccharides including immunomodulatory, antitumor, antimicrobial effects and hypocholesterolemic effects [11–13]. With excellent properties (including highly stable, safe, nontoxic, hydrophilic and biodegradable) and abundant resources, polysaccharides have been studied and applied in biomaterial fields. Particularly, the application of polysaccharides in nanoparticle delivery systems (NPDSs) has attracted increasing attention in recent decades. Nanoparticles are defined as solid, colloidal particles consisting of macromolecules with a size in the range of 10–1000 nm [14]. Nanoparticle material is an important application field of nanotechnology, which has been applied in various fields such as food, feed, biomedical sciences, and drug/gene delivery systems [15–17]. In particular, NPDSs have been attracting increasing attention recently and are became a focus. NPDSs are generally referred to as nanometric carriers with various morphologies, including nanospheres, nanocapsules, nanomicelles, nanoliposomes, and nanodrugs, etc. [18]. Presently, nanoparticles have been widely employed to deliver food ingredients, drugs, polypeptides, proteins, genes, and other biomolecules.

8.2 Nanoparticles for delivery systems Previous studies and applications of nanoparticles were mainly focused on drug delivery, because nanoparticles can entrap drugs or biomolecules into their interior structures and/or absorb drugs or biomolecules onto their exterior surfaces. Therefore, the most important application of nanoparticles was nanovectors. Additionally, previous studies have summarized various outstanding advantages of nanoparticle drug delivery systems, including [19]: (1) they can pass through the smallest capillary vessels because of their ultra-tiny volume and avoid rapid clearance by phagocytes so that their duration in blood stream is greatly prolonged; (2) they can penetrate cells and tissue gaps to arrive at target organs such as liver, spleen, lung, spinal cord and lymph; (3) they could show controlled release properties due to the biodegradability, pH, ion and/or temperature sensibility of materials; (4) they can improve the utility of drugs and reduce toxic side effects; etc. With these superiorities, nanoparticle drug delivery systems have been widely studied in biological, medical and pharmaceutical applications. Meanwhile, nanoparticles have also attracted increasing attention for food delivery systems to provide maximum protection for sensitive food components against oxidation, enzyme degradation and pH before reaching the target [20, 21]. Several bioactive food components have been found effective in treatment of coronary heart diseases, inflammation, and immune disorder etc. in corporation with the diet [21, 22]. They are mainly grouped into isoprenoids, fatty acids, proteins and amino acids, polysaccharides and minerals.

8.3 Polysaccharides and their nanoparticles

| 161

However, some of these components have poor properties, such as instability during digestion, poor solubility and bioavailability, ingredient interactions, and unpleasant taste. Thereby, nanoparticles were employed to improve the bioavailability of bioactive food ingredients, provide maximum stability, introduce controlled/target release of encapsulated compound during mastication and digestion for efficient absorption into the body system. Based on the method of preparation, nanoparticles can be designed and constructed to possess different properties and release characteristics for the best delivery or encapsulation of the therapeutic agent [23]. The nanoparticles currently used and studied as nanovectors can be grouped into three main classes or “generations” [24, 25]. The first class focuses on a passive delivery system for the target site. For example, the size of particles could enable the driving systems to the tumor site, but not specific recognition of the targets [26]. The second class of nanovectors includes additional functional groups that allow for molecular recognition of the target tissue. These functional groups include ligands, aptamers, and small peptides that bind to specific target-cell surface markers or surface markers expressed in the disease microenvironment [27]. Meanwhile, pH-sensitive polymers are included in this category. Finally, the third class aimed to successfully overcome the natural barriers that the vector needs to efficiently deliver the drug to the target site. This goal will only be reached by a “multistage” approach, and such a system has been recently reported [28]. Currently, particles of the first generation have been approved by FDA for their use in metastatic breast cancer [29]. Numerous clinical trials are also ongoing for the targeted second class nanovectors, particularly in cancer applications [30]. Compared with nondegradable materials, biodegradable systems have some advantages in the application of nanoparticles, including nontoxic, biotolerable, biocompatible, biodegradable, and water-soluble properties. Polysaccharides, as the most popular natural biopolymer, have their unique features in developing nanoparticles [31–33].

8.3 Polysaccharides and their nanoparticles A variety of polysaccharides have been modified with various reactants and investigated for the synthesis and application of nanoparticles using various methods [34–36]. From the viewpoint of polyelectrolyte, polysaccharides can be divided into nonpolyelectrolytes (including starch, dextran, cellulose, etc.) and polyelectrolytes as listed in Tab. 8.1, the latter can be further divided into positively charged polysaccharides (chitosan) and negatively charged polysaccharides (alginate, pectin, hyaluronic acid, etc.).

Cell wall of green plants, many forms of algae and the oomycetes Product of many bacterial strains and enzymatic product of cell-free culture supernatant Enzymatic degradation of starch derived from potatoes, corn, rice and other sources

Cell wall of all plants Vertebrate organisms

104 –106 Da

107 –108 Da

2 × 103 –107 Da

From thousands to 2 000 000 Da

NA

3800–20 000 Da

200 000–500 000 Da

50 000–180 000 Da

Can reach as high as 107 Da

Cellulose

Dextran

Cyclodextrin

Pullulan

Guar gum

Chitosan

Alginate

Pectin

Hyaluronic acid

Extraction from cell walls and intercellular spaces of marine brown algae

Shells of crab, shrimp and krill; cell walls of fungi

Extraction of the seeds of Cyamopsis tetragonoloba

Bacterial homopolysaccharide production from starch by the fungus Aureobasidium pullulans

Higher plants

Source

Amylose: Da Amylopectin: 106– 107 Da

107 –108

Molecular weight (Da)

Starch

Polysaccharides

Tab. 8.1: Polysaccharides in the preparation of nanoparticle delivery systems.

Nanoparticles for wound healing and skin regeneration [44], for siRNA delivery [63], anticancer drug delivery [64], etc.

Facilitation of the delivery of specific sequences of amino acids [60], drugs [61]; wound healing scaffolds [62], etc.

Nanocarrier gene delivery [56], for protein delivery [57], for drug delivery [58]; scaffolds in tissue engineering [59], etc.

Nanocarrier in drug delivery system [54], nanocontainer for hydrophobic drugs [55], nanoparticles complex with other molecules as discussed below, etc.

Nanocomposite for drug release [53]

Drug and gene delivery [49], protein delivery [50], tissue engineering [51], wound healing [52], etc.

Nanocarrier for drug release control [47], for gene and drug delivery [48], etc.

Controlled release of growth factors for wound healing and skin regeneration [44], drug nanocarrier [45], delivery for delicate bioactive molecules [46], etc.

Nanodelivery for hydrophobic active ingredients [41], for oral protein [42], for anticancer drugs [43], etc.

Nanocarrier for parenteral drug delivery [37], for colon-specific delivery [38], for insulin controlled release [39], for siRNA delivery [40], etc.

Application in nanoparticles

162 | 8 Novel nanoparticle materials for drug/food delivery-polysaccharides

8.3 Polysaccharides and their nanoparticles

| 163

8.3.1 Nonpolyelectrolyte polysaccharides 8.3.1.1 Starch and its derivation Starch-based nanoparticles have attracted increasing attention due to their good hydrophilicity, biocompatibility and biodegradability. Starch is made up of two main structural components: amylose and amylopectin [65, 66], the former consists of a linear backbone of α-1,4-linked glucose with/without a low level of branching with a α-1,6-linkage, while amylopectin is a highly branched form of ‘amylose’ [39, 67] (Fig. 8.1). As the second most abundant biomaterial in nature, starch has been modified with various reactants by way of chemical reaction with hydroxyl groups in the starch molecule for preparation of nanoparticles [68]. OH O HO O HO

HO HO

Amylose

HO

O OH

O HO

HO

O OH

O HO n

HO O

O OH

O OH

OH O O

HO

Amylopectin

OH O

HO

HO H OH O HO

O OH O

Fig. 8.1: Chemical structure of starch (adapted from [2]).

The hydrophobic derivative of starch by grafting hydrophobic poly (lactic acid) chains (PLA) was prepared to nanoparticles through crosslinked method and used for drug delivery taking Indomethacin as the model drug [69, 70]. Besides, propyl-starch nanoparticles were prepared to entrap docetaxel for cancer therapy and revealed high encapsulation efficiency [71, 72].

8.3.1.2 Cellulose and its derivation Cellulose is the most abundant polysaccharide available on Earth with the formula of (C6 H10 O5 )n [73–75]. The cellulose molecule is formed by a linear chain of β-1,4-linked D-glucose units with different lengths (Fig. 8.2). Due to the insolubility of cellulose, nanoparticles are usually prepared from its derivative. Several studies have been conducted on the Poly (ε-caprolactone) (PCL) and poly (L-lactic acid) (PLLA) modification of soluble cellulose and its derivate [76–78]. Besides, chitosan or its oligomer could complex carboxymethyl cellulose (CMC, anionic derivative of cellulose) to form stable cationic nanoparticles for coating with plasmid DNA in genetic immunization [79].

164 | 8 Novel nanoparticle materials for drug/food delivery-polysaccharides

OH *

O

O

O

HO

OH

HO

* O

OH OH

Fig. 8.2: Chemical structure of cellulose (adapted from [2]).

n

8.3.1.3 Dextran and its derivation Dextrans are a class of polysaccharides consisting of a linear backbone with mainly α1,6-linked glucose, and a variable amount of α (1 → 2), α (1 → 3) and α (1 → 4) branched linkages [80] (Fig. 8.3). Currently, dextran is widely applied in fields of chemical, pharmaceutical, clinical and food industry playing the function of adjuvant, emulsifier, carrier, drug, stabilizer, and thickener of jam and ice cream [81, 82]. Dextrans are colloidal, hydrophilic and water-soluble substances, and can be decomposed in human feces due to bacterial action [44]. Hence, various drug-dextran prodrugs could be able to keep integrity in stomach and the small intestine but release in colon [83].

O O OH O

O OH alfa–1,6 alfa–1,4

O n

OH OH OH alfa–1,6

m

Fig. 8.3: Chemical structure of dextran (adapted from [2]).

Dextrans have been modified to extend their surface-active properties and potential applications in pharmacy, biochemistry and medicine. Either water-soluble or waterinsoluble dextran derivatives have been prepared based on the extent of modification. The water-insoluble derivatives could be solubilized in organic solvents like tetrahydrofuran or dichloromethane saturated with water [84–86]. With food availability, biocompatibility and biodegradability, dextran has been widely selected as promising biomaterial in the preparation of nanovectors.

8.3.1.4 Cyclodextrins and their derivation Cyclodextrins (CDs) are cyclic oligosaccharides composed of at least five α-1,4-linked glucopyranose units in a rigid 4 C1 chair conformation, prepared by enzymatic degradation of starch. The most common CDs contain 6–8 D-glucose units and are known

| 165

8.3 Polysaccharides and their nanoparticles

OH

O

HO

O

OHOHO

HO O OH

α OH O OH O HO

HO

OH

OH HO

O OH O HO

OH O

OH O OH

OH HO OH O O

O

O

O OH HO

O OH O

O HO OH

O HO

OH O OH O

OH OHO O

HO OH O

OH O

HO

OH O OH

γ

OH

OH O OH O

OH

OH O

O

OH O OH

OH O HO

HO

OH O OH

β

O

O OH HO

O HO OH

O

OH

HO

HO

O

HO

HO O OH O OH HO O

O

OH

Fig. 8.4: Chemical structure of cyclodextrins.

as α-CD, β-CD, and γ-CD, respectively, as shown in Fig. 8.4 [87–89]. CDs are neither hydrolyzed nor absorbed in stomach and small intestine, but are absorbed in the large intestine so that the vast microflora present in the colon could degrade them into small saccharides [90, 91]. This property ensures CDs as a colon targeting carrier. Cyclodextrins are considered to be the most widely explored materials applied for nanoparticle formation. CD-based nanoparticles were usually prepared by selfassembling method, because the three-dimensional ring structure of CDs allows for encapsulation of hydrophobic molecules within the oligosaccharide cavity [92– 94]. Harada et al. have reported that αCD-based nanoparticles could be formed with poly(ethylene glycol) (PEG) for drug delivery [95]. Besides, modified CDs have been used to prepare nanoparticles for delivery of small interfering RNA (siRNA), as well as controlled release of antimalarial artemisinin [96].

8.3.1.5 Pullulan Pullulan is a linear glucan produced from starch by the fungus Aureobasidium pullulans [97, 98]. The backbone of pullulan is formed by maltotriose units (α-1,4-Dglucopyranose) through α-1,6 glycosidic linkage in a ratio of 2 : 1 (Fig. 8.5). Due to existence of an α-1,6 linkage in the molecule, pullulan tends to perform as a random flexible coil in aqueous solution, which may result in its biodegradability and it has high adhesion, structural flexibility and solubility [99]. The FDA has approved pullulan for various applications, due to its hemocompatible, nonimmunogenic, and noncarcinogenic properties [100]. The applications of pullulan extend to a variety of fields: in biomedical fields as drug and gene delivery [101], tissue engineering [102], and wound healing [103]; in pharmaceuticals as a coating agent [49, 52]; in foods and beverages as a filler; as an edible, mostly tasteless polymer, as well as edible films [104, 105].

166 | 8 Novel nanoparticle materials for drug/food delivery-polysaccharides

O HO

CH2

HO

O

O

OH

O

OH

OH

OH O

O OH

OH

O OH

CH2

Fig. 8.5: Chemical structure of pullulan (adapted from [2]).

Pullulan needs to be modified with hydrophobic molecules for self-assembling in water solution which will then behave as carriers of agents. Hydrophobic molecules including cholesterol, hexadecanol, vitamin H, etc. have been used to derive pullulan to obtain amphiphilic micelles [19]. The partially hydrophobized pullulan shows unique association and potential application in delivery systems. For example, both cholesterol-pullulan and a copolymer of N-isopropylacrylamide and N-[4(1-pyrenyl)butyl]-N-noctadecylacrylamide, and hexadecyl group-bearing pullulan have been self-assembly prepared for nanoparticle delivery carriers [106–108]. Pullulan acetate (PA) is the other important hydrophobized pullulan, which can form self-aggregation nanoparticles as well as its modified materials [109, 110].

8.3.1.6 Guar gum Guar gum, also called guaran, is formed by a linear chain of β-1,4-D-mannopyranosyl residues with branching points at O-6 site having α-D-galactopyranosyl units as the side chains (Fig. 8.6) [111]. With water solubility, guar gum is a nonionic natural polysaccharide derived from the seeds of Cyamopsistetra gonolobusis. Guar gum hydrates in cold water to form highly viscous colloidal dispersions or sols [112]. Guar gum solution is stable under pH range 5–7, but extreme pH and high temperature conditions (e.g. pH 3 at 50 °C) can degrade its structure [113]. With properties of being nontoxic, highly viscous and easily available, guar gum is commonly used for various applications: in food industry as thickener for sauces, ice creams, etc.; in pharmaceuticals as binder and disintegrant for solid dosage and as hydrophilic matrix for oral controlled release dosage [111, 112]. Besides, guar gum has been extensively applied in colon-specific drug delivery due to its drug sustained release property and susceptibility to microbial degradation in the colon [114]. In addition, guar gum has been found to be a better stabilizer of the nanoparticles [113].

8.3 Polysaccharides and their nanoparticles

| 167

CH2OH O

OH OH

O OH

CH2 O

O OH

OH

OH

OH

OH

O

OH

CH2OH n Fig. 8.6: Chemical structure of guar gum.

8.3.2 Positively charged polyelectrolyte polysaccharides 8.3.2.1 Chitin and chitosan Chitin is the main component of fungal cell walls, the exoskeleton of crustaceans (such as crabs and shrimp) and insects [115–117]. As shown in Fig. 8.7, chitin consists of a linear chain of N-acetylglucosamine. The role of chitin is analogous to cellulose in structure and to keratin in function. As the deacetylation product of chitin, chitosan is composed of glucosamine and N-acetylglucosamine by β-1,4-glycosidic bonds forming linear backbone. Chitosan is produced industrially by alkali treatment to hydrolyze the amino-acetyl groups of chitin with the degree of deacetylation ( %DD) in range 60–100 % [117, 118]. CH3

OH O O

O HO

NH O

OH NH

HO O

HO HO

O

OH

O NH2

O

HO

OH

O NH2

OH CH3

O

HO n

O NH2

OH

n

Chitin

Chitosan

Fig. 8.7: Chemical structure of chitin and chitosan.

Chitosan is considered as a biocompatible, biodegradable and nontoxic biomaterial and widely applied in pharmaceutical and biomedical fields [119]. In the field of nanomedicine, chitosan has received considerable attention as vector in novel bioadhesive drug delivery systems which prolong the residence time of the drugs at the site of absorption and increase the drug bioavailability [54]. However, this biopolymer can only solubilize in diluted acidic aqueous solution (pH < 6.5) for the glucosamine units converting into a soluble form with protonated amine groups [120]. The insol-

168 | 8 Novel nanoparticle materials for drug/food delivery-polysaccharides

ubility in water and organic solvents limited its application, but chitosan could be hydrophobically modified to obtain nanoparticles and applied as nanocarriers for drugs due to their biocompatibility in vivo [121]. Experimental in vitro and in vivo results show chitosan as a promising nanocarrier for controlled release of various drugs with excellent encapsulated efficacy.

8.3.3 Negatively charged polyelectrolyte polysaccharides 8.3.3.1 Alginate Alginate is an anionic linear polysaccharide derived from cell walls and intercellular spaces of marine brown algae. The structure of alginate consists of a backbone of β-1,4-linked D-mannuronic acid (M unit) and α-1,4-linked L-guluronic acid (G unit) arranged of various compositions and sequences depending on the source of the alginate (Fig. 8.8). M block segments provide linear and flexible conformation, while G block segments serve folded and rigid structural conformations [122]. Moreover, the ratio of M unit against G unit has been reported to affect its physicochemical properties, as well as its further applications [100].

COOH O OH

O OH OH COOH O O

OH

O m

n

Fig. 8.8: Chemical structure of alginate.

Alginate is a biopolymer with biocompatible, nonimmunogenic, nontoxic and biodegradable properties [123]. A large number of free hydroxyl and carboxyl groups in the backbone of alginate may be modified to achieve solubility, hydrophobicity, physicochemical and biological characteristics, as well as various potential applications [122]. The applications of alginate have extended to various industries: as food additive and thickener in food industry [124]; as scaffolds in tissue engineering [125]; and as controlled drug release devices in biomedicine [126]. Besides, previous studies have indicated that the muco-adhesive, biocompatible and biodegradable properties of alginate make it an important and hopeful tool in the preparation of controlled drug-delivery systems achieving an enhanced drug bioavailability [124, 126].

8.3 Polysaccharides and their nanoparticles

| 169

8.3.3.2 Pectin Pectin exists in the cell wall of plants to function as cell adhesion. Pectins are a family of complex polysaccharides containing 1,4-linked D-galacturonic acid residues and were usually divided into homogalacturonans (HG), rhamnogalacturonan I (RG-I), and rhamnogalacturonan II (RG-II), xylogalacturonan (XGA) and apiogalacturonan (AGA) based on their structural features [61, 127]. The common pectin type is HG as shown in Fig. 8.9. COOCH3

COOH

O

O O

OH

O

OH

OH OH

Fig. 8.9: Chemical structure of pectin.

Pectin can resist the degradation in the physiological environment of the stomach and the small intestine, but can be decomposed by pectinases secreted by microflora of the human colon [128]. Thanks to these properties, pectin could function as prebiotics and delivery vector for components from the mouth to the colon [129, 130]. However, pectin cannot protect its encapsulated components during its delivery through the stomach and small intestine due to its high water solubility [129, 130]. Hence, studies mainly focused on pectin derivatives with water resistant and enzymatic degradation. For this purpose, calcium pectinate was deeply studied as a drug carrier for colonspecific delivery because this complex can reduce the solubility of pectin and keep stable in low pH environment [128]. Besides, pectin has been combined with other polymers, including 4-aminothiophenol [131], chitosan [132], hyaluronic acid [133] or poly (lactide-co-glycolide) [134], showing good results as controlled drug release devices.

8.3.3.3 Hyaluronic acid Hyaluronic acid (HA) (also called hyaluronan, hyaluronate) is an anionic polysaccharide composed of repeating disaccharide units of D-glucuronic acid and N-acetyl D-glucosamine linked via altering β-1,3 and β-1,4 glycosidic bonds as shown in Fig. 8.10 [135]. HA has been reported to associate with several cellular processes, including angiogenesis and the regulation of inflammation [136]. As described in previous studies, HA is a biodegradable, bioactive, nonimmunogenic and noncytotoxic polysaccharide [137]. Similar to other glycosaminoglycans, hyaluronan can function as a targeting vector for the delivery of chemotherapeutic agents to cancerous tissues, as many tumors overexpress the hyaluronan CD44 and

170 | 8 Novel nanoparticle materials for drug/food delivery-polysaccharides CH2OH O O

COO O

OH O

NHCOCH3

OH n OH

Fig. 8.10: Repeating unit of hyaluronic acid.

RHAMM receptors [138]. As a drug delivery carrier, HA has several advantages including the negligible nonspecific interaction with serum components due to its polyanionic characteristics [139] and the highly efficient targeted specific delivery to the liver tissues with HA receptors [140].

8.3.4 Hyperbranched polysaccharides The above described polysaccharides are mainly attributed to linear polysaccharides or partially branched polysaccharides. However, hyperbranched polysaccharides (HBPs), including amylopectin, glycogen and some glucans from mushroom cell walls, have attracted increasing attention in the fields of nanotechnology and pharmacology because of their unique structures and properties [141–143]. The spherical architecture of highly branched macromolecules provides numerous terminal units that can be converted into various functional groups leading to novel nanomaterials [32, 144]. HBPs can also form polymeric micelles that are spherical aggregates of amphiphilic blocks of copolymers which enhance water solubility and decrease the toxicity of hydrophobic drugs [145]. Compared with synthetic highly branched polymers, research interest in natural HBPs is emerging in the field of biomaterials due to their nontoxicity, good biocompatibility and biodegradability [19]. Unlike other highly branched polymers, spherical HBPs do not only provide reaction sites for the formation of nanoparticles, but they also protect the nanoparticles in a shell structure with excellent dispersion in water [32, 33, 146]. These unique properties of HBPs can be applied in the fields of drug delivery and controlled release [33, 146]. For example, hyperbranched cationic amylopectin derivatives have been designed for gene delivery with high transfection efficiency and exhibited potential as nonviral gene vectors [147]. HBPs were shown to interact strongly with lectins due to the clustering or multivalent effects of the numerous nonreducing saccharide units on their surfaces [148]. HBPs have also potent bioactivity including immune-modulatory and antitumor effects [31, 149]. Recent studies on HBPs have been focused on obtaining them from natural sources and characterizing their physical properties including solubility, shrinking factors, and rheological properties [150–152].

8.4 Nanoparticle preparation based on polysaccharides

|

171

8.3.5 Other polysaccharides Besides, various fungal polysaccharides have attracted increasing attention in the dispersion of nanoparticles. These polysaccharides mainly contain helical chains structures, including lentinan, scleroglucan, schizophyllan, etc. Lentinan is a structural polysaccharide from the fruiting body of Lentinus edodes, with a β-(1→3)-Dglucan backbone and two β-(1→6) linked glucoses as side groups every five (1→3)-Dglucoside residues. Lentinan has been identified as triple helical chains in aqueous solutions and single flexible chains in DMSO or high concentration of alkali solutions. These structural properties of lentinan have been applied to disperse silver nanoparticles in water [153]. Triple helical schizophyllan has also been applied as reducing and stabilizing agent to prepare silver nanoparticles [154]. Moreover, CaCO3–lentinan microspheres have been obtained by self-assembly nanoparticles and applied as an anticancer drug carrier [155]. Scleroglucan (SCL) is the other β-(1→3) and β-(1→6) glucan with triple helical chains, produced by fungi of the genus Sclerotium. SCL was used to prepare scleroglucan gels for investigation of drug-loading effects on release by using theophylline as the model drug [156]. In addition, SCL-PVA (polyvinylalcohol) hydrogels containing magnetic nanoparticles have been prepared to study drug release behavior [157].

8.4 Nanoparticle preparation based on polysaccharides Recent reviews have presented excellent summaries of the preparation and application of polysaccharide-based nanoparticles, mainly focusing on starch, chitosan, cellulose, pectin, etc. Along with the study of nanoparticles furthering the ‘third generation’, more polysaccharide-based nanovectors emerged in the field of drug, gene and food delivery systems. Based on structural characteristics, polysaccharide nanoparticles are prepared mainly by four mechanisms, including covalent crosslinking, ionic crosslinking, polyelectrolyte complexation, and self-assembly of hydrophobically modified polysaccharides.

8.4.1 Covalent crosslinking polysaccharide nanoparticles The method of covalent crosslinking was first performed on the preparation of chitosan-based nanoparticles. In this method, a crosslinker is necessary to obtain the desired nanoparticles. For example, glutaraldehyde was used as the crosslinker to crosslink chitosan-based nanoparticles [158, 159]. However, the toxicity of glutaraldehyde on cell viability limits its utility in delivery systems. Carbodiimide was then employed as a water-soluble condensation agent and biocompatible crosslinker for covalent crosslinking [160]. With the aid of this crosslinker, natural di- and tri-

172 | 8 Novel nanoparticle materials for drug/food delivery-polysaccharides

carboxylic acids were used for intermolecular crosslinking of chitosan nanoparticles [160, 161]. Furthermore, hyaluronic acid has also been used to prepare nanoparticles by using a carbodiimide method [162]. The nanoparticles prepared with this method were mainly in the form of polycations, polyanions, and polyampholytes and stable in aqueous media at low pH, neutral, and mild alkaline conditions. In the swollen state, the average size of the particles was in the range of 270–370 nm depending on the pH.

8.4.2 Ionic crosslinking polysaccharide nanoparticles Similar to the method of covalent crosslinking, ionic crosslinking aims at charged polysaccharides or modified polysaccharides. However, ionic crosslinking has unique advantages against covalent crosslinking, including mild preparation conditions and simple procedures. For ionic polysaccharides, low MW of polyanions and polycations could act as ionic crosslinkers for polycationic and polyanionic polysaccharides, respectively. To date, the most widely used polyanion crosslinker is tripolyphosphate (TPP), which was first used to crosslink chitosan nanoparticles in 1997 [163, 164]. TPP is nontoxic and has multivalent anions. It can form a gel by ionic interaction between positively charged amino groups of chitosan and negatively charged counterions of TPP [165]. Currently, chitosan is usually replaced by water-soluble chitosan derivatives to prepare nanoparticles by ionic crosslinking method. Compared with chitosan itself, its derivatives can easily dissolve in neutral aqueous media, avoiding the potential toxicity of acids and hence protecting the bioactivity of loaded biomacromolecules. For instance, N-trimethyl chitosan nanoparticles have been synthesized by ionic crosslinking of N-trimethyl chitosan with TPP and showed an encapsulation efficiency up to 95 % and a loading capacity up to 50 % (w/w) [159, 166]. Their potential as a carrier system was evaluated for the nasal delivery of proteins, ovalbumin [167]. The following studies indicated the nontoxicity of N-trimethylchitosan/TPP nanoparticles to Calu-3 cells. The absorption properties of N-trimethylchitosan/TPP nanoparticles have also been evaluated by use of in vitro (Caco-2 cells) and ex vivo (excised rat jejunum) models [168]. Besides, some negatively charged polysaccharides (including alginate, pectin and hyaluronic acid) bearing carboxylic groups can be crosslinked by bivalent calcium ion to form nanoparticles. For example, Ca-alginate nanoparticles have been prepared by water-in-oil reverse microemulsion method (~ 80 nm in size) [169] and ion-induced gelification method (235.5 nm in size) [170] to deliver gene and drugs, respectively. The relative bioavailabilities of all drugs encapsulated were significantly higher than oral free drugs. In drug delivery system, all drugs (isoniazid, pyrazinamide, rifampicin) were detected in organs (lungs, liver and spleen) above the minimum inhibitory concentration until 15 days post nebulization, whilst free drugs stayed up to day 1. These

8.4 Nanoparticle preparation based on polysaccharides

| 173

inhalable nanoparticles could serve as an ideal carrier for the controlled release of antitubercular drugs.

8.4.3 Polyelectrolyte complexing polysaccharide nanoparticles Polysaccharide nanoparticles by polyelectrolyte complexation (PEC) are also based on polysaccharides or their derivatives with charge. PEC has received increasing attention compared with other nanoparticle preparation techniques, because PEC-based nanoparticles have several characteristics favorable for cellular uptake and colloidal stability, including suitable diameter and surface charge, spherical morphology and a low polydispersity index (PdI), and so on [171]. Besides, nanoparticles prepared by this method not only avoid many kinds of aggression in harsh conditions (such as organic solvents and sonication during preparation), but also keep the stability and biological activity of the encapsulated agents in completely aqueous condition and in ambient temperature [172, 173].

Electrostastic interaction

+

Polymers with positive charge

Polymers or Drugs

Polyelectrolyte complexes

Fig. 8.11: The schematic illustration of the nanoparticles formed by polyelectrolyte complexation [174].

As shown in Fig. 8.11, polysaccharides can form PEC with oppositely charged polymers by intermolecular electrostatic interaction. Currently, chitosan is the only natural polycationic polysaccharide to form nanoparticles in this method, for its watersoluble and biocompatible properties [175]. Chitosan-based PEC nanoparticles can be synthesized with many negative polymers, including polysaccharides [176], peptides [177], polyacrylic acid family [178], and so on [179, 180].

8.4.4 Self-assembly polysaccharide nanoparticles Self-assembling polysaccharide nanoparticles have moved to the forefront due to their unique advantages, including biocompatibility and stimulus responsiveness. The selfassembled nanoparticles are composed of a core of hydrophobic moieties surrounding by a hydrophilic outer shell, which could serve as protection for the carried hydropho-

174 | 8 Novel nanoparticle materials for drug/food delivery-polysaccharides

bic agents [181–183]. By grafting with hydrophobic segments, polysaccharides can form amphiphilically polymeric micelles which enhance water solubility and decrease the toxicity of hydrophobic drugs [145]. In aqueous environment, polyamphiphiles spontaneously form micelles or micelle-like aggregates via undergoing intra- or intermolecular associations between hydrophobic moieties, in order to minimize interfacial free energy. In recent years, numerous studies have been carried out to investigate the synthesis and the application of polysaccharide-based self-aggregate nanoparticles as drug delivery systems. Generally, these hydrophobic molecules can be divided into linear [184, 185], cyclic hydrophobic molecules [186, 187], polyacrylate family [188, 189], etc.

8.5 Applications of polysaccharide-based nanoparticles In the design of nanoscale carriers, polysaccharides have received considerable attention for their unique properties, including safety, nontoxicity, bioavailability and biocompatibility. Currently, polysaccharide-based nanoparticles have been investigated and applied in medical and food fields as new promising biomaterials.

8.5.1 Medical applications 8.5.1.1 Delivery of peptides and proteins In recent years, some peptides and proteins have been discovered for therapeutic and antigenic bioactivities and attracted considerable attention [190]. However, most of these biologically derived drugs are limited for in vivo application by their disadvantages like low stability, short biological half-life and the need to cross biological barriers. From this viewpoint, polysaccharide-based nanoparticles can overcome some of the problems of systemic administration. Currently, polysaccharide-based nanoparticles have been prepared and applied for delivery of several protein drugs, including insulin [191, 192], basic fibroblast growth factor (bFGF) [193, 194] and epidermal growth factor receptor (EGFR) antisense (AS) [195]. Meanwhile, bovine serum albumin (BSA) is a normal model in the preparation of nanoparticles based on polysaccharides [196]. Insulin is one of the most widely applied therapeutic peptides for the treatment of insulin-dependent diabetes mellitus. The normal problems of oral insulin administration are low bioavailability on acidic gastric pH, the enzymatic barrier of the intestinal tract and the physical barrier made up of the intestinal epithelium. Therefore, polysaccharide-based nanoparticles have been prepared for the inclusion of insulin in various studies by different methods, which show good release control and good results in loading efficacy [192]. Nanoparticles based on alginate-alginate coated with chitosan, alginate-dextran have been prepared by the interaction of carboxylic groups of alginate with amino groups of insulin [197, 198]. Insulin-loaded nanoparticles have

8.5 Applications of polysaccharide-based nanoparticles

|

175

also been synthesized by ionic crosslinked method using chitosan as the selected polysaccharide and TPP anions as crosslinker agent [199]. Additionally, PEC between chitosan and different polyanions (alginate, glucomannan, and dextran sulfate) has been prepared for insulin inclusion in some studies and shown ~90 % association efficiency values. Self-assembling method was also used to prepare insulin-loaded nanoparticles using cholesterol-bearing pullulan as the selected polysaccharide [106]. These insulin-loaded polysaccharide-based nanoparticles showed excellent behavior in evaluation of physiological activity.

8.5.1.2 Delivery of anticancer drugs The main problems of cancer chemotherapy are related with the toxicity caused by anticancer drugs on normal tissues and release control of drugs on the target site [200]. Thus, nanoparticles are being extensively investigated as carriers of anticancer drugs, in order to overcome several problems in cancer chemotherapy, including reducing harmful side effects, enhancing blood circulation time, controlling the release concentration of the drug at the tumor site for a desired time period, thus, increasing therapeutic efficiency. Among the available potential drug carrier systems in nanoscale, polysaccharide-based nanoparticles play an important role and their use with some anticancer drugs shows promising results [201]. Tamoxifen, a drug for hormone dependent breast cancer, has been entrapped into polysaccharide-based nanoparticles to overcome the undesirable side effects and to increase the concentration at the tumor site due to specific recognition for targeting tissue or organ. Tamoxifen-loaded nanoparticles were prepared by Sarmah and coworkers based on guar gum, which is commonly used for colon specific drug delivery in the pharmaceutical industry [111, 202]. As a breast cancer drug, mitoxantrone is positively charged and then has been encapsulated in chitosan-based nanoparticles by ion gelation method using sodium TPP as gelation agent, and obtaining an encapsulation efficacy of 98 % [203]. There were also other anticancer drugs delivered by polysaccharide-based nanoparticles, including methotrexate (MTX, a folate antimetabolite) [204], doxorubicin (DOX, an anthracycline ring antibiotic drug) [205], paclitaxel (an anticancer drug) [206], etc.

8.5.1.3 Nanovectors of nucleic acids and genetic material Up to now, gene therapy has been applied in many different diseases such as cancer, AIDS, and cardiovascular diseases [207]. Gene therapy aims to transfer genetic materials into specific cells of a patient to repair defective genes responsible for disease development [208]. To transfer the genes to the specific site, genes must escape the processes that affect the disposition of macromolecules and avoid the degradation by serum nuclease.

176 | 8 Novel nanoparticle materials for drug/food delivery-polysaccharides

Small interfering RNAs (siRNAs) have been employed as a novel tool to block the expression of infectious diseases and cancers. However, siRNA suffers particular problems including poor cellular uptake, rapid degradation as well as limited blood stability. For this reason, chitosan-based nanoparticles have been prepared to transfect small interfering RNAs (siRNAs) by modified ionic gelation method with TPP as crosslinker agent [209]. Besides, PEC between chitosan and different polyanions has been used to prepare nanoparticles in order to include nucleic acids, since chitosanDNA nanoparticles demonstrated low transfection efficiencies and the incorporation of secondary polymers improved the characteristics of these systems [210]. Recently, a new method has been used to prepare a gene nanocarrier based on triple helical β-glucan [211]. The target DNA sequence was firstly bound to polydeoxyadenylic acid (poly (dA)) by disulfide bonds (poly (dA)–SS–DNA). Lentinan was then used to combine the poly (dA)–SS–DNA chain to form a new triple helical conformation and provide protection of the delivered DNA. The target DNA was then delivered into the cell via endocytosis and released from the delivery system by automatic cleavage disulfide bonds in cytoplasm.

8.5.2 Food applications Apart from delivery of drugs and genes, polysaccharide-based nanoparticles have also been studied and applied in delivery systems of food bioactives. Antioxidants, probiotics, polyunsaturated fatty acids, and proteins are common bioactives that can be added to food to improve nutritional value, to prevent diseases and to improve overall health [212]. Nanodelivery of these components may improve their stability [213, 214], solubility [215, 216], functionality [217, 218], cellular uptake [219–221], and bioavailability [222–224] and may also provide controlled release [225–227] for better efficacy of the bioactive. Nanoparticles formed by polysaccharides can deliver a variety of lipophilic bioactives to the colon, maintain their integrity and are kept impermeable within the upper gastrointestinal tract (GIT). It may be necessary to design a nanoparticle or microgel that protects the bioactive component within a food product, but that can release it within the upper GIT so that it can be absorbed. For instance, casein-pectin microgels have been reported to encapsulate polyunsaturated lipids and protect them from oxidation [228], but they will fully dissociate under simulated GIT conditions [229, 230]. In this case, microgel dissociation takes place due to weakening of the electrostatic forces holding them together, as well as digestion of the casein molecules by proteases. Polysaccharides have also been used as stabilizing agents to stabilize emulsions for providing controlled release, improving entrapment efficiency, and protection from degradation [231, 232]. In this case, the hydrophobic food bioactive is to be dissolved in the internal organic phase of an oil-in-water emulsion, whereas double

8.6 Conclusions

|

177

emulsions are employed for nanodelivery of hydrophilic molecules [233, 234]. Besides, gum arabic maltodextrin was developed to improve the stability and bioavailability of epigallocatechin gallate [235].

8.6 Conclusions As reviewed above, so many polysaccharides and their derivatives are employed as one of the most used biomaterials in preparation of nanoparticulate delivery systems. Due to their complex structure, polysaccharides show variability and versatility, which is difficult to reproduce with synthetic polymers. A variety of polysaccharidebased nanoparticles have been obtained by various preparation methods towards three evolution aspects: need for less toxic agents, simplification of the procedures and optimization to improve yield and entrapment efficiency. Now it is possible to choose the best method of preparation and the best suitable polymer to achieve an efficient encapsulation of the drug/gene/food ingredient, taking into account the agent features in this selection. In addition, a variety of novel polysaccharides with specific properties, such as hyperbranched polysaccharides, are being selected to prepared nanoparticles for delivery systems. Until now, these nanoparticles have been investigated in terms of their physicochemical properties, drug-loading efficiency, in vitro toxicity, and comparatively simple in vivo tests. Deeper studies, such as the specific interaction of these nanoparticles with human organs, tissues, cells, or biomolecules, as well as how the administration of these systems can affect the metabolism, need to be carried out and focused on in the future. A combination of in silico, in vitro, and in vivo studies is required for the safe application of nanoparticle delivery systems in drugs, genes and food.

References [1] [2] [3] [4] [5] [6] [7]

Ren L, Perera C, Hemar Y. Antitumor activity of mushroom polysaccharides: A review, Food and Function, 2012, 3(11), 1118–30. Namazi H, Fathi F, Heydari A. Nanoparticles based on modified polysaccharides, in Hashim AA, ed. The delivery of nanoparticles, InTech, 2012, pp. 149–84. Aminabhavi TM, Balundgi RH, Cassidy PE. A review on biodegradable plastics, PolymerPlastics Technology and Engineering, 1990, 29(3), 235–62. Van Soest PV, Robertson JB, Lewis BA. Methods for dietary fiber, neutral detergent fiber, and nonstarch polysaccharides in relation to animal nutrition, J Dairy Sci, 1991, 74(10), 3583–97. Hoover R. Composition, molecular structure, and physicochemical properties of tuber and root starches: A review, Carbohyd Polym, 2001, 45(3), 253–67. Chadha MJ. Novel techniques for the characterisation of exopolysaccharides secreted by lactic acid bacteria. 2009, University of Huddersfield: Huddersfield. Meléndez R, Meléndez-Hevia E, Canela EI. The fractal structure of glycogen: A clever solution to optimize cell metabolism, Biophys J, 1999, 77(3), 1327–32.

178 | 8 Novel nanoparticle materials for drug/food delivery-polysaccharides

[8] [9] [10] [11]

[12]

[13] [14] [15] [16] [17] [18] [19] [20] [21]

[22] [23] [24]

[25] [26] [27]

[28]

[29]

Astbury WT, Davies MM. Structure of cellulose, Nature, 1944, 154, 84. Dweltz NE. The structure of chitin, Biochimica et biophysica acta, 1960, 44, 416–35. Zhou JS, Preparation of mushroom-source chitin from mushroom root, Peop. Rep. China: CN, 2010, p. 4p. Zhang M, Cheung PCK, Chiu LCM, Wong EYL, Ooi VEC. Cell-cycle arrest and apoptosis induction in human breast carcinoma Mcf-7 cells by carboxymethylated β-glucan from the mushroom sclerotia of Pleurotus tuber-regium, Carbohyd Polym, 2006, 66(4), 455–62. Lin Y, Zhang L, Chen L, Jin Y, Zeng F, Jin J, Wan B, Cheung PCK. Molecular mass and antitumor activities of sulfated derivatives of α-glucan from Poria Cocos mycelia, Int J Biol Macromol, 2004, 34(5), 231–36. Wasser SP. Medicinal mushrooms as a source of antitumor and immunomodulating polysaccharides, Appl Microbiol Biot, 2002, 60(3), 258–74. Hamidi M, Azadi A, Rafiei P. Hydrogel nanoparticles in drug delivery, Adv Drug Deliver Rev, 2008, 60(15), 1638–49. Colvin VL, Schlamp MC, Alivisatos AP. Light-emitting diodes made from cadmium selenide nanocrystals and a semiconducting polymer, Nature, 1994, 370(6488), 354–57. Dickinson E. Use of nanoparticles and microparticles in the formation and stabilization of food emulsions, Trends Food Sci Tech, 2012, 24(1), 4–12. Nitta S, Numata K. Biopolymer-based nanoparticles for drug/gene delivery and tissue engineering, Int J Mol Sci, 2013, 14(1), 1629–54. Acosta E. Bioavailability of nanoparticles in nutrient and nutraceutical delivery, Curr Opin Colloid In, 2009, 14(1), 3–15. Liu Z, Jiao Y, Wang Y, Zhou C, Zhang Z. Polysaccharides-based nanoparticles as drug delivery systems, Adv Drug Deliver Rev, 2008, 60(15), 1650–62. Adjonu R, Doran G, Torley P, Agboola S. Whey protein peptides as components of nanoemulsions: a review of emulsifying and biological functionalities, J Food Eng, 2014, 122, 15–27. Borel T, Sabliov CM. Nanodelivery of bioactive components for food applications: types of delivery systems, properties, and their effect on adme profiles and toxicity of nanoparticles, Annu Rev Food Sci Technol, 2014, 5, 197–213. McClements DJ. Nanoscale nutrient delivery systems for food applications: improving bioactive dispersibility, stability, and bioavailability, J Food Sci, 2015, 80(7), N1602–11. Barratt GM. Therapeutic applications of colloidal drug carriers, Pharmaceutical Science & Technology Today, 2000, 3(5), 163–71. Martínez A, Fernández A, Pérez E, Benito M, Teijón JM, Blanco MD. Polysaccharide-based nanoparticles for controlled release formulations, in Hashim A, ed. The delivery of nanoparticles, InTech, 2012. Sakamoto J, Annapragada A, Decuzzi P, Ferrari M. Antibiological barrier nanovector technology for cancer applications, Expert Opin Drug Deliv, 2007, 4(4), 359–69. Romberg B, Hennink WE, Storm G. Sheddable coatings for long-circulating nanoparticles, PHARM Res-DORDR, 2008, 25(1), 55–71. Kang J, Lee MS, Copland Iii JA, Luxon BA, Gorenstein DG. Combinatorial selection of a single stranded DNA thioaptamer targeting Tgf-B1 protein, Bioorg Med Chem Lett, 2008, 18(6), 1835–39. Tasciotti E, Liu X, Bhavane R, Plant K, Leonard AD, Price BK, Cheng MM, Decuzzi P, Tour JM, Robertson F, Ferrari M. Mesoporous silicon particles as a multistage delivery system for imaging and therapeutic applications, Nat Nanotechnol, 2008, 3(3), 151–57. Kratz F. Albumin as a drug carrier: design of prodrugs, drug conjugates and nanoparticles, J Control Release, 2008, 132(3), 171–83.

References

[30] [31]

[32] [33]

[34] [35] [36] [37] [38]

[39]

[40]

[41] [42] [43]

[44]

[45]

[46]

[47]

| 179

Gomes BAR, Moreira IES, Rocha S, Coelho M, Pereira MDC. Polysaccharide-based nanoparticles for cancer therapy, Journal of Nanopharmaceutics and Drug Delivery, 2013, 1(4), 335–54. Kuang HX, Xia YG, Liang J, Yang BY, Wang QH, Wang XG. Structural characteristics of a hyperbranched acidic polysaccharide from the stems of Ephedra sinica and its effect on T-cell subsets and their cytokines in Dth mice, Carbohyd Polym, 2011, 86(4), 1705–11. Zhang YF, Structure, chain conformation and functional modification of hyperbranched polysaccharide. Wuhan University: Wuhan, China, 2011, pp. 16–37. Satoh T. Synthesis and encapsulation-release property of unimolecular inversed micelle having hyperbranched polysaccharide core, in. Callaos N, et al., eds. WMSCI, 2006: 10th World Multi-Conference On Systemics, Cybernetics And Informatics, 2006, pp. 15–16. Namazi H, Mosadegh M. Preparation and properties of starch/nanosilicate layer/polycaprolactone composites, J Polym Environ, 2011, 19(4), 980–87. Namazi H, Dadkhah A. Convenient method for preparation of hydrophobically modified starch nanocrystals with using fatty acids, Carbohyd Polym, 2010, 79(3), 731–37. Aumelas A, Serrero A, Durand A, Dellacherie E, Leonard M. Nanoparticles of hydrophobically modified dextrans as potential drug carrier systems, Colloid Surface B, 2007, 59(1), 74–80. Narayanan D, Nair S, Menon D. A systematic evaluation of hydroxyethyl starch as a potential nanocarrier for parenteral drug delivery, Int J Biol Macromol, 2015, 74, 575–84. Sivapragasam N, Thavarajah P, Ohm J, Ohm J, Margaret K, Thavarajah D. Novel starch based nano scale enteric coatings from soybean meal for colon-specific delivery, Carbohyd Polym, 2014, 111, 273–79. Zhang Z, Shan H, Chen L, He C, Zhuang X, Chen X. Synthesis of ph-responsive starch nanoparticles grafted poly (L-glutamic acid) for insulin controlled release, Eur Polym J, 2013, 49(8), 2082–91. Amar-Lewis E, Azagury A, Chintakunta R, Goldbart R, Traitel T, Prestwood J, Landesman-Milo D, Peer D, Kost J. Quaternized starch-based carrier for sirna delivery: from cellular uptake to gene silencing, J Control Release, 2014, 185, 109–20. Numata Y, Mazzarino L, Borsali R. A slow-release system of bacterial cellulose gel and nanoparticles for hydrophobic active ingredients, Int J Pharmaceut, 2015, 486(1–2), 217–25. Song Y, Chen L. Effect of net surface charge on physical properties of the cellulose nanoparticles and their efficacy for oral protein delivery, Carbohyd Polym, 2015, 121, 10–17. Elumalai R, Patil S, Maliyakkal N, Rangarajan A, Kondaiah P, Raichur AM. Protaminecarboxymethyl cellulose magnetic nanocapsules for enhanced delivery of anticancer drugs against drug resistant cancers, Nanomedicine: Nanotechnology, Biology and Medicine, 2015, 11(4), 969–81. Gainza G, Villullas S, Pedraz JL, Hernandez RM, Igartua M. Advances in drug delivery systems (DDSs) to release growth factors for wound healing and skin regeneration, Nanomedicine: Nanotechnology, Biology and Medicine, 2015, 11(6), 1551–73. Saboktakin MR, Tabatabaie RM, Maharramov A, Ramazanov MA. Synthesis and characterization of ph-dependent glycol chitosan and dextran sulfate nanoparticles for effective brain cancer treatment, Int J Biol Macromol, 2011, 49(4), 747–51. Parraga JE, Zorzi GK, Diebold Y, Seijo BA, Sanchez A. Nanoparticles based on naturallyoccurring biopolymers as versatile delivery platforms for delicate bioactive molecules: an application for ocular gene silencing, Int J Pharmaceut, 2014, 477(1–2), 12–20. Anirudhan TS, Divya PL, Nima J. Synthesis and characterization of silane coated magnetic nanoparticles/glycidylmethacrylate-grafted-maleated cyclodextrin composite hydrogel as a drug carrier for the controlled delivery of 5-fluorouracil, Materials Science and Engineering: C, 2015, 55, 471–81.

180 | 8 Novel nanoparticle materials for drug/food delivery-polysaccharides

[48]

[49] [50]

[51] [52] [53] [54] [55]

[56]

[57]

[58] [59]

[60] [61] [62]

[63]

[64]

[65] [66]

Kang L, Gao Z, Huang W, Jin M, Wang Q. Nanocarrier-mediated co-delivery of chemotherapeutic drugs and gene agents for cancer treatment, Acta Pharmaceutica Sinica B, 2015, 5(3), 169–75. Singh RS, Kaur N, Kennedy JF. Pullulan and pullulan derivatives as promising biomolecules for drug and gene targeting, Carbohyd Polym, 2015, 123, 190–207. Dionísio M, Cordeiro C, Remu Án-López C, Seijo BA, Rosa Da Costa AM, Grenha A. Pullulanbased nanoparticles as carriers for transmucosal protein delivery, Eur J Pharm Sci, 2013, 50(1), 102–13. Lamichhane A, Azegami T, Kiyono H. The mucosal immune system for vaccine development, Vaccine, 2014, 32(49), 6711–23. Prajapati VD, Jani GK, Khanda SM. Pullulan: An exopolysaccharide and its various applications, Carbohyd Polym, 2013, 95(1), 540–49. Islan GA, Mukherjee A, Castro GR. Development of biopolymer nanocomposite for silver nanoparticles and ciprofloxacin controlled release, Int J Biol Macromol, 2015, 72, 740–50. Varum FJ, McConnell EL, Sousa JJ, Veiga F, Basit AW. Mucoadhesion and the gastrointestinal tract, Critical Reviews™in Therapeutic Drug Carrier Systems, 2008, 25(3). Opanasopit P, Ngawhirunpat T, Chaidedgumjorn A, Rojanarata T, Apirakaramwong A, Phongying S, Choochottiros C, Chirachanchai S. Incorporation of camptothecin into N-phthaloyl chitosan-g-mpeg self-assembly micellar system, Eur J Pharm Biopharm, 2006, 64(3), 269–76. Jain S, Tran T, Amiji M. Macrophage repolarization with targeted alginate nanoparticles containing Il-10 plasmid DNA for the treatment of experimental arthritis, Biomaterials, 2015, 61, 162–77. Mukhopadhyay P, Chakraborty S, Bhattacharya S, Mishra R, Kundu PP. Ph-Sensitive chitosan/alginate core-shell nanoparticles for efficient and safe oral insulin delivery, Int J Biol Macromol, 2015, 72, 640–48. Bhujbal SV, de Vos P, Niclou SP. Drug and cell encapsulation: alternative delivery options for the treatment of malignant brain tumors, Adv Drug Deliver Rev, 2014, 67–68, 142–53. Kim H, Jung G, Yoon J, Han J, Park Y, Kim D, Zhang M, Kim D. Preparation and characterization of nano-sized hydroxyapatite/alginate/chitosan composite scaffolds for bone tissue engineering, Materials Science and Engineering: C, 2015, 54, 20–25. Ugurlu T, Turkoglu M, Gurer US, Akarsu BG. Colonic delivery of compression coated nisin tablets using pectin/hpmc polymer mixture, Eur J Pharm Biopharm, 2007, 67(1), 202–10. Zhang W, Xu P, Zhang H. Pectin in cancer therapy: a review, Trends Food Sci Tech, 2015, 44(2), 258–71. Ninan N, Muthiah M, Park I, Kalarikkal N, Elain A, Wui Wong T, Thomas S, Grohens Y. Wound healing analysis of pectin/carboxymethyl cellulose/microfibrillated cellulose based composite scaffolds, Mater Lett, 2014, 132, 34–37. Lee MS, Lee JE, Byun E, Kim NW, Lee K, Lee H, Sim SJ, Lee DS, Jeong JH. Target-specific delivery of sirna by stabilized calcium phosphate nanoparticles using dopa–hyaluronic acid conjugate, J Control Release, 2014, 192, 122–30. Han HS, Choi KY, Ko H, Jeon J, Saravanakumar G, Suh YD, Lee DS, Park JH. Bioreducible corecrosslinked hyaluronic acid micelle for targeted cancer therapy, J Control Release, 2015, 200, 158–66. Rodrigues A, Emeje M. Recent applications of starch derivatives in nanodrug delivery, Carbohyd Polym, 2012, 87(2), 987–94. Dufresne A. Crystalline starch based nanoparticles, Curr Opin Colloid In, 2014, 19(5), 397–408.

References

[67]

[68] [69] [70] [71]

[72] [73] [74] [75] [76] [77] [78]

[79] [80] [81] [82] [83] [84]

[85]

[86]

[87]

|

181

Namazi H, Dadkhah A. Surface modification of starch nanocrystals through ring-opening polymerization of epsilon-caprolactone and investigation of their microstructures, J Appl Polym Sci, 2008, 110(4), 2405–12. Le Corre D, Bras J, Dufresne A. Starch nanoparticles: A review, Biomacromolecules, 2010, 11(5), 1139–53. Simi CK, Abraham TE. Hydrophobic grafted and cross-linked starch nanoparticles for drug delivery, Bioproc Biosyst Eng, 2007, 30(3), 173–80. Thielemans W, Belgacem MN, Dufresne A. Starch nanocrystals with large chain surface modifications, Langmuir, 2006, 22(10), 4804–10. Santander-Ortega MJ, Stauner T, Loretz B, Ortega-Vinuesa JL, Bastos-Gonz A Lez D, Wenz G, Schaefer UF, Lehr CM. Nanoparticles made from novel starch derivatives for transdermal drug delivery, J Control Release, 2010, 141(1), 85–92. Dandekar P, Jain R, Stauner T, Loretz B, Koch M, Wenz G, Lehr C. A hydrophobic starch polymer for nanoparticle-mediated delivery of docetaxel, Macromol Biosci, 2012, 12(2), 184–94. Mohanty AK, Misra M, Hinrichsen G. Biofibres, Biodegradable polymers and biocomposites: an overview, Macromol Mater Eng, 2000, 276(1), 1–24. Riedel U, Nickel JOR. Natural fibre-reinforced biopolymers as construction materials – new discoveries, Die Angewandte Makromolekulare Chemie, 1999, 272(1), 34–40. Bledzki AK, Gassan J. Composites reinforced with cellulose based fibres, Prog Polym Sci, 1999, 24(2), 221–74. Teramoto Y, Nishio Y. Cellulose diacetate-graft-poly(lactic acid)s: synthesis of wide-ranging compositions and their thermal and mechanical properties, Polymer, 2003, 44(9), 2701–09. Shi RW, Burt HM. Synthesis and characterization of amphiphilic hydroxypropylcellulose-graftpoly(epsilon-caprolactone), J Appl Polym Sci, 2003, 89(3), 718–27. Teramoto Y, Yoshioka M, Shiraishi N, Nishio Y. Plasticization of cellulose diacetate by graft copolymerization of epsilon-caprolactone and lactic acid, J Appl Polym Sci, 2002, 84(14), 2621–28. Cui Z, Mumper RJ. Chitosan-based nanoparticles for topical genetic immunization, J Control Release, 2001, 75(3), 409–19. Misaki A, Torii M, Sawai T, Goldstein IJ. Structure of the dextran of leuconostoc mesenteroides B-1355, Carbohyd Res, 1980, 84(2), 273–85. Prado HJ, Matulewicz MC. Cationization of polysaccharides: A path to greener derivatives with many industrial applications, Eur Polym J, 2014, 52, 53–75. McCurdy RD, Goff HD, Stanley DW, Stone AP. Rheological properties of dextran related to food applications, Food Hydrocolloid, 1994, 8(6), 609–23. Shukla RK, Tiwari A. Carbohydrate Polymers: Applications and recent advances in delivering drugs to the colon, Carbohyd Polym, 2012, 88(2), 399–416. Bertholon I, Vauthier C, Labarre D. Complement activation by core-shell poly (isobutylcyanoacrylate)-polysaccharide nanoparticles: influences of surface morphology, length, and type of polysaccharide, Pharm Res-Dordr, 2006, 23(6), 1313–23. Durand A, Marie E, Rotureau E, Leonard MEL, Dellacherie E. Amphiphilic polysaccharides: useful tools for the preparation of nanoparticles with controlled surface characteristics, Langmuir, 2004, 20(16), 6956–63. Rouzes C, Leonard M, Durand A, Dellacherie E. Influence of polymeric surfactants on the properties of drug-loaded Pla nanospheres, Colloids and Surfaces B: Biointerfaces, 2003, 32(2), 125–35. Larsen KL, Endo T, Ueda H, Zimmermann W. Inclusion complex formation constants of alpha-, beta-, gamma-, delta-, epsilon-, zeta-, eta- and theta-cyclodextrins determined with capillary zone electrophoresis, Carbohyd Res, 1998, 309(2), 153–59.

182 | 8 Novel nanoparticle materials for drug/food delivery-polysaccharides

[88] [89] [90] [91] [92] [93]

[94] [95] [96] [97] [98] [99] [100] [101] [102]

[103]

[104]

[105]

[106]

[107]

Miyazawa I, Ueda H, Nagase H, Endo T, Kobayashi S, Nagai T. Physicochemical properties and inclusion complex-formation of delta-cyclodextrin, Eur J Pharm Sci, 1995, 3(3), 153–62. Fujiwara T, Tanaka N, Kobayashi S. Structure of delta-cyclodextrin 13.75H2O, Chem Lett, 1990, 5, 739–42. Uekama K, Hirayama F, Irie T. Cyclodextrin drug carrier systems, Chem Rev, 1998, 98(5), 2045–76. Connors KA. The stability of cyclodextrin complexes in solution, Chem Rev, 1997, 97(5), 1325–58. Uekama K. Recent aspects of pharmaceutical application of cyclodextrins, J Incl Phenom Macro, 2002, 44(1–4), 3–07. Daoud-Mahammed S, Couvreur P, Bouchemal K, Ch E Ron M, Lebas GEV, Amiel C, Gref R. Cyclodextrin and polysaccharide-based nanogels: entrapment of two hydrophobic molecules, benzophenone and tamoxifen, Biomacromolecules, 2009, 10(3), 547–54. Liu Y, Zhao Y, Zhang H. Recognition-induced supramolecular porous nanosphere formation from cyclodextrin conjugated by cholic acid, Langmuir, 2006, 22(7), 3434–38. Harada A, Kamachi M. Complex formation between poly (ethylene glycol) and α-cyclodextrin, Macromolecules, 1990, 23(10), 2821–23. Davis ME. The first targeted delivery of sirna in humans via a self-assembling, cyclodextrin polymer-based nanoparticle: from concept to clinic, Mol Pharmaceut, 2009, 6(3), 659–68. Glinel K, Huguet J, Muller G. Comparison of the associating behaviour between neutral and anionic alkylperfluorinated pullulan derivatives, Polymer, 1999, 40(25), 7071–81. Bataille I, Huguet J, Muller G, Mocanu G, Carpov A. Associative behaviour of hydrophobically modified carboxymethylpullulan derivatives, Int J Biol Macromol, 1997, 20(3), 179–91. Leathers TD. Biotechnological production and applications of pullulan, Appl Microbiol Biot, 2003, 62(5-6), 468–73. Coviello T, Matricardi P, Marianecci C, Alhaique F. Polysaccharide hydrogels for modified release formulations, J Control Release, 2007, 119(1), 5–24. Rekha MRCP. Pullulan as a promising biomaterial for biomedical applications: a perspective, Trends in Biomaterial & Artificial Organs, 2007, 116. Thebaud NEL, Pierron DEE, Bareille R, Le Visage C, Letourneur D, Bordenave L. Human endothelial progenitor cell attachment to polysaccharide-based hydrogels: a pre-requisite for vascular tissue engineering, Journal of Materials Science: Materials in Medicine, 2007, 18(2), 339–45. Bae H, Ahari AF, Shin H, Nichol JW, Hutson CB, Masaeli M, Kim S, Aubin H, Yamanlar S, Khademhosseini A. Cell-laden microengineered pullulan methacrylate hydrogels promote cell proliferation and 3D cluster formation, Soft Matter, 2011, 7(5), 1903–11. Khanzadi M, Jafari SM, Mirzaei H, Chegini FK, Maghsoudlou Y, Dehnad D. Physical and mechanical properties in biodegradable films of whey protein concentrate–pullulan by application of beeswax, Carbohyd Polym, 2015, 118, 24–29. Synowiec A, Gniewosz MG, Kra Niewska K, Przyby JAL, B Czek K, W Glarz Z. Antimicrobial and antioxidant properties of pullulan film containing sweet basil extract and an evaluation of coating effectiveness in the prolongation of the shelf life of apples stored in refrigeration conditions, Innov Food Sci Emerg, 2014, 23, 171–81. Akiyoshi K, Kobayashi S, Shichibe S, Mix D, Baudys M, Kim SW, Sunamoto J. Self-assembled hydrogel nanoparticle of cholesterol-bearing pullulan as a carrier of protein drugs: complexation and stabilization of insulin, J Control Release, 1998, 54(3), 313–20. Akiyoshi K, Deguchi S, Moriguchi N, Yamaguchi S, Sunamoto J. Self-aggregates of hydrophobized polysaccharides in water. Formation and characteristics of nanoparticles, Macromolecules, 1993, 26(12), 3062–68.

References

|

183

[108] Jung SW, Jeong YL, Kim YH, Kim SH. Self-assembled polymeric nanoparticles of poly(ethylene glycol) grafted pullulan acetate as a novel drug carrier, Arch Pharm Res, 2004, 27(5), 562–69. [109] Zhang HZ, Gao FP, Liu LR, Li XM, Zhou ZM, Yang XD, Zhang QQ. Pullulan acetate nanoparticles prepared by solvent diffusion method for epirubicin chemotherapy, Colloids & Surfaces B Biointerfaces, 2009, 71(1), 19–26. [110] Park KH, Song HC, Na K, Bom HS, Lee KH, Kim S, Kang D, Lee DH. Ionic strength-sensitive pullulan acetate nanoparticles (pan) for intratumoral administration of radioisotope: ionic strength-dependent aggregation behavior and (99m)technetium retention property, Colloids Surf B Biointerfaces, 2007, 59(1), 16–23. [111] Sarmah JK, Rita M, Saibal Kanti B, Ranadeep M, Angshuman B. Controlled release of tamoxifen citrate encapsulated in cross-linked guar gum nanoparticles, Int J Biol Macromol, 2011, 49(3), 390–96. [112] Barbucci R, Pasqui D, Favaloro R, Panariello G. A thixotropic hydrogel from chemically crosslinked guar gum: synthesis, characterization and rheological behaviour, Carbohyd Res, 2008, 343(18), 3058–65. [113] Tiraferri A, Kai LC, Sethi R, Elimelech M. Reduced aggregation and sedimentation of zerovalent iron nanoparticles in the presence of guar gum, Journal of Colloid & Interface Science, 2008, 324(1–2), 71–79. [114] Soumya RS, Ghosh S, Abraham ET. Preparation and characterization of guar gum nanoparticles, Int J Biol Macromol, 2010, 46(2), 267–69. [115] Kean T, Thanou M, Roth S. Trimethylated chitosans as non-viral gene delivery vectors: cytotoxicity and transfection efficiency, J Control Release, 2005, 103(103), 643–53. [116] Ramesh HP, Tharanathan RN. Carbohydrates – the renewable raw materials of high biotechnological value, Crit Rev Biotechnol, 2003, 23(2), 149–73. [117] Yuan Z. Study on the synthesis and catalyst oxidation properties of chitosan bound nickel(II) complexes, Chemical Industry Times, 2007. [118] Muzzarelli RAA, Muzzarelli C. Chitosan chemistry: relevance to the biomedical sciences, Polysaccharides I, 2005, 151–209. [119] Guerrero S, Teijón C, Mu Iz E, Teijón JM, Blanco MD. Characterization and in vivo evaluation of ketotifen-loaded chitosan microspheres, Carbohyd Polym, 2010, 79(4), 1006–13. [120] Sinha VR, Singla AK, Wadhawan S, Kaushik R, Kumria R, Bansal K, Dhawan S. Chitosan microspheres as a potential carrier for drugs, Int J Pharm, 2004, 274(1–2), 1–33. [121] Yoo HS, Lee JE, Chung H, Kwon IC, Jeong SY. Self-assembled nanoparticles containing hydrophobically modified glycol chitosan for gene delivery, J Control Release, 2005, 103(1), 235–43. [122] Yang J, Xie Y, He W. Research progress on chemical modification of alginate: a review, Carbohyd Polym, 2011, 84(1), 33–39. [123] Rinaudo M. Biomaterials based on a natural polysaccharide: alginate, Tip, 2014, 17(1), 92–96. [124] Nair LS, Laurencin CT. Biodegradable polymers as biomaterials, Prog Polym Sci, 2007, 32(8–9), 762–98. [125] Barbosa MA, Granja PL, Barrias CC, Amaral IF. Polysaccharides as scaffolds for bone regeneration, Itbm-Rbm, 2005, 26(3), 212–17. [126] Pandey R, Ahmad Z. Nanomedicine and experimental tuberculosis: facts, flaws, and future, Nanomedicine: Nanotechnology, Biology and Medicine, 2011, 7(3), 259–72. [127] Posé S, Kirby AR, Paniagua C, Waldron KW, Morris VJ, Quesada MA, Mercado JA. The nanostructural characterization of strawberry pectins in pectate lyase or polygalacturonase silenced fruits elucidates their role in softening, Carbohyd Polym, 2015, 132, 134–45. [128] Liu L, Fishman ML, Kost J, Hicks KB. Pectin-based systems for colon-specific drug delivery via oral route, Biomaterials, 2003, 24(19), 3333–43.

184 | 8 Novel nanoparticle materials for drug/food delivery-polysaccharides

[129] Yang L, Chu JS, Fix JA. Colon-specific drug delivery: new approaches and in vitro/in vivo evaluation, Int J Pharmaceut, 2002, 235(1–2), 1–15. [130] Sinha VR, Kumria R. Polysaccharides in colon-specific drug delivery, Int J Pharmaceut, 2001, 224(1–2), 19–38. [131] Perera G, Barthelmes J, Bernkop-Schn U Rch A. Novel pectin-4-aminothiophenole conjugate microparticles for colon-specific drug delivery, J Control Release, 2010, 145(3), 240–46. [132] Fernandez-Hervas MJ, Fell JT. Pectin/chitosan mixtures as coatings for colon-specific drug delivery: an in vitro evaluation, Int J Pharmaceut, 1998, 169(1), 115–19. [133] Pliszczak DEE, Bourgeois S, Bordes C, Valour J, Mazoyer MEE, Orecchioni AM, Nakache E, Lant E, Ri P. Improvement of an encapsulation process for the preparation of pro-and prebioticsloaded bioadhesive microparticles by using experimental design, Eur J Pharm Sci, 2011, 44(1), 83–92. [134] Liu L, Won YJ, Cooke PH, Coffin DR, Fishman ML, Hicks KB, Ma PX. Pectin/poly (lactide-coglycolide) composite matrices for biomedical applications, Biomaterials, 2004, 25(16), 3201–10. [135] Cafaggi S, Russo E, Stefani R, Parodi B, Caviglioli G, Sillo G, Bisio A, Aiello C, Viale M. Preparation, characterisation and preliminary antitumour activity evaluation of a novel nanoparticulate system based on a cisplatin-hyaluronate complex and N-trimethyl chitosan, Invest New Drug, 2011, 29(3), 443–55. [136] Leach JB, Schmidt CE. Characterization of protein release from photocrosslinkable hyaluronic acid-polyethylene glycol hydrogel tissue engineering scaffolds, Biomaterials, 2005, 26(2), 125–35. [137] Oh EJ, Park K, Kim KS, Kim J, Yang J, Kong J, Lee MY, Hoffman AS, Hahn SK. Target specific and long-acting delivery of protein, peptide, and nucleotide therapeutics using hyaluronic acid derivatives, J Control Release, 2010, 141(1), 2–12. [138] Yip GW, Smollich M, Götte M. Therapeutic value of glycosaminoglycans in cancer, Mol Cancer Ther, 2006, 5(9), 2139–48. [139] Ito T, Iida-Tanaka N, Niidome T, Kawano T, Kubo K, Yoshikawa K, Sato T, Yang Z, Koyama Y. Hyaluronic acid and its derivative as a multi-functional gene expression enhancer: protection from non-specific interactions, adhesion to targeted cells, and transcriptional activation, J Control Release, 2006, 112(3), 382–88. [140] Zhou B, McGary CT, Weigel JA, Saxena A, Weigel PH. Purification and molecular identification of the human hyaluronan receptor for endocytosis, Glycobiology, 2003, 13(5), 339–49. [141] Satoh T, Kakuchi T. Synthesis of hyperbranched carbohydrate polymers by ring-opening multibranching polymerization of anhydro sugar, Macromol Biosci, 2007, 7(8), 999–1009. [142] Satoh T, Ishihara H, Maeda T, Kaga H, Kakuchi T. Synthesis of hyperbranched polysaccharide by thermally induced cationic polymerization of 1,6-anhydro sugar, Abstracts of Papers of the American Chemical Society, 2002, 224, U359. [143] Kadokawa J, Tagaya H. Architecture of polysaccharides with specific structures: synthesis of hyperbranched polysaccharides, Polym Advan Technol, 2000, 11(3), 122–26. [144] Tao YZ, Zhang LN, Yan F, Wu XJ. Chain conformation of water-insoluble hyperbranched polysaccharide from fungus, Biomacromolecules, 2007, 8(7), 2321–28. [145] Kitajyo Y, Imai T, Sakai Y, Tamaki M, Tani H, Takahashi K, Narumi A, Kaga H, Kaneko N, Satoh T, Kakuchi T. Encapsulation-release property of amphiphilic hyperbranched D-glucan as a unimolecular reverse micelle, Polymer, 2007, 48(5), 1237–44. [146] Kitajyo Y, Sakai Y, Imai T, Satoh T, Kaga H, Kakuchi T. Capsulation-release property of amphiphilic hyperbranched polysaccharide for hydrophilic guest molecules, Abstracts of Papers of the American Chemical Society, 2004, 227, U372–73.

References

|

185

[147] Zhou YF, Yang B, Ren XY, Liu ZZ, Deng Z, Chen LM, Deng YB, Zhang LM, Yang LQ. Hyperbranched cationic amylopectin derivatives for gene delivery, Biomaterials, 2012, 33(18), 4731–40. [148] Hoai NT, Sasaki A, Sasaki M, Kaga H, Kakuchi T, Satoh T. Synthesis, characterization, and lectin recognition of hyperbranched polysaccharide obtained from 1,6-anhydro-Dhexofuranose, Biomacromolecules, 2011, 12(5), 1891–99. [149] Huang ZP, Zhang LN, Duan XB, Liao ZQ, Ding H, Cheung PCK. Novel highly branched watersoluble heteropolysaccharides as immunopotentiators to inhibit s-180 tumor cell growth in Balb/C mice, Carbohyd Polym, 2012, 87(1), 427–34. [150] Tao YZ, Yan Y, Xu WL. Shrinking factors of hyperbranched polysaccharide from fungus, Carbohyd Res, 2009, 344(11), 1311–18. [151] Tao YZ, Xu WL. Microwave-assisted solubilization and solution properties of hyperbranched polysaccharide, Carbohyd Res, 2008, 343(18), 3071–78. [152] Tao YZ, Zhang LN. Determination of molecular size and shape of hyperbranched polysaccharide in solution, Biopolymers, 2006, 83(4), 414–23. [153] Li S, Zhang Y, Xu X, Zhang L. Triple helical polysaccharide-induced good dispersion of silver nanoparticles in water, Biomacromolecules, 2011, 12(8), 2864–71. [154] Abdel-Mohsen AM, Abdel-Rahman RM, Fouda MMG, Vojtova L, Uhrova L, Hassan AF, Al-Deyab SS, El-Shamy IE, Jancar J. Preparation, characterization and cytotoxicity of schizophyllan/silver nanoparticle composite, Carbohyd Polym, 2014, 102, 238–45. [155] Ma X, Yuan S, Yang L, Li L, Zhang X, Su C, Wang K. Fabrication and potential applications of CaCO3 -lentinan hybrid materials with hierarchical composite pore structure obtained by selfassembly of nanoparticles, Crystengcomm, 2013, 15(41), 8288–99. [156] François NJ, Rojas AM, Daraio ME. Rheological and drug-release behaviour of a scleroglucan gel matrix at different drug loadings, Polym Int, 2005, 54(12), 1613–19. [157] François NJ, Allo S, Jacobo SE, Daraio ME. Composites of polymeric gels and magnetic nanoparticles: preparation and drug release behavior, J Appl Polym Sci, 2007, 105(2), 647–55. [158] Kadam AA, Lee DS. Glutaraldehyde cross-linked magnetic chitosan nanocomposites: reduction precipitation synthesis, characterization, and application for removal of hazardous textile dyes, Bioresource Technol, 2015, 193, 563–67. [159] Anitha A, Sowmya S, Kumar PTS, Deepthi S, Chennazhi KP, Ehrlich H, Tsurkan M, Jayakumar R. Chitin and chitosan in selected biomedical applications, Prog Polym Sci, 2014, 39(9), 1644–67. [160] Yin L, Ding J, He C, Cui L, Tang C, Yin C. Drug permeability and mucoadhesion properties of thiolated trimethyl chitosan nanoparticles in oral insulin delivery, Biomaterials, 2009, 30(29), 5691–700. [161] Bodn A R M, Hartmann JF, BORBELY J. Nanoparticles from chitosan, Polymer Preprints, American Chemical Society, Division of Polymer Chemistry, 2004, 45(2), 307–08. [162] Saravanakumar G, Choi KY, Yoon HY, Kim K, Park JH, Kwon IC, Park K. Hydrotropic hyaluronic acid conjugates: synthesis, characterization, and implications as a carrier of paclitaxel, Int J Pharmaceut, 2010, 394(1–2), 154–61. [163] Calvo P, Remunan-Lopez C, Vila-Jato JL, Alonso MJ. Novel hydrophilic chitosan-polyethylene oxide nanoparticles as protein carriers, J Appl Polym Sci, 1997, 63(1), 125–32. [164] Calvo P, Remu N An-L O Pez C, Vila-Jato JL, Alonso MIAJ. Chitosan and chitosan/ethylene oxide-propylene oxide block copolymer nanoparticles as novel carriers for proteins and vaccines, Pharm Res-Dordr, 1997, 14(10), 1431–36. [165] Jain D, Banerjee R. Comparison of ciprofloxacin hydrochloride-loaded protein, lipid, and chitosan nanoparticles for drug delivery, Journal of Biomedical Materials Research Part B: Applied Biomaterials, 2008, 86(1), 105–12.

186 | 8 Novel nanoparticle materials for drug/food delivery-polysaccharides

[166] Martins AF, Facchi SP, Monteiro JP, Nocchi SR, Silva CTP, Nakamura CV, Girotto EM, Rubira AF, Muniz EC. Preparation and cytotoxicity of N,N,N-trimethyl chitosan/alginate beads containing gold nanoparticles, Int J Biol Macromol, 2015, 72, 466–71. [167] Amidi M, Romeijn SG, Borchard G, Junginger HE, Hennink WE, Jiskoot W. Preparation and characterization of protein-loaded N-trimethyl chitosan nanoparticles as nasal delivery system, J Control Release, 2006, 111(1), 107–16. [168] Sandri G, Bonferoni MC, Rossi S, Ferrari F, Gibin S, Zambito Y, Di Colo G, Caramella C. Nanoparticles Based On N-trimethylchitosan: evaluation of absorption properties using in vitro (caco-2 cells) and ex vivo (excised rat jejunum) models, Eur J Pharm Biopharm, 2007, 65(1), 68–77. [169] You J, Peng C. Calcium-alginate nanoparticles formed by reverse microemulsion as gene carriers, Wiley Online Library, 2005, pp. 147–53. [170] Zahoor A, Sharma S, Khuller GK. Inhalable alginate nanoparticles as antitubercular drug carriers against experimental tuberculosis, Int J Antimicrob Ag, 2005, 26(4), 298–303. [171] Boddohi S, Moore N, Johnson PA, Kipper MJ. Polysaccharide-based polyelectrolyte complex nanoparticles from chitosan, heparin, and hyaluronan, Biomacromolecules, 2009, 10(6), 1402–09. [172] Luo Y, Wang Q. Recent development of chitosan-based polyelectrolyte complexes with natural polysaccharides for drug delivery, Int J Biol Macromol, 2014, 64, 353–67. [173] Hou R, Nie L, Du G, Xiong X, Fu J. Natural polysaccharides promote chondrocyte adhesion and proliferation on magnetic nanoparticle/PVA composite hydrogels, Colloids and Surfaces B: Biointerfaces, 2015, 132, 146–54. [174] Hu Y, Yang T, Hu X. Novel polysaccharides-based nanoparticle carriers prepared by polyelectrolyte complexation for protein drug delivery, Polym Bull, 2012, 68(4), 1183–99. [175] Silva DA, Maciel JS, Feitosa J, Paula H, De Paula R. Polysaccharide-based nanoparticles formation by polyeletrolyte complexation of carboxymethylated cashew gum and chitosan, J Mater Sci, 2010, 45(20), 5605–10. [176] Li T, Shi X, Du Y, Tang Y. Quaternized chitosan/alginate nanoparticles for protein delivery, J Biomed Mater Res A, 2007, 83(2), 383–90. [177] Lee P, Peng S, Su C, Mi F, Chen H, Wei M, Lin H, Sung H. The use of biodegradable polymeric nanoparticles in combination with a low-pressure gene gun for transdermal DNA delivery, Biomaterials, 2008, 29(6), 742–51. [178] Sajeesh S, Sharma CP. Novel pH responsive polymethacrylic acid-chitosan-polyethylene glycol nanoparticles for oral peptide delivery, Journal of Biomedical Materials Research Part B: Applied Biomaterials, 2006, 76(2), 298–305. [179] Jayakumar R, Chennazhi KP, Muzzarelli RAA, Tamura H, Nair SV, Selvamurugan N. Chitosan conjugated DNA nanoparticles in gene therapy, Carbohyd Polym, 2010, 79(1), 1–8. [180] Guo C, Gemeinhart RA. Understanding the adsorption mechanism of chitosan onto poly(lactide-co-glycolide) particles, Eur J Pharm Biopharm, 2008, 70(2), 597–604. [181] Lee KY, Jo WH, Kwon IC, Kim Y, Jeong SY. Structural determination and interior polarity of selfaggregates prepared from deoxycholic acid-modified chitosan in water, Macromolecules, 1998, 31(2), 378–83. [182] Nishikawa T, Akiyoshi K, Sunamoto J. Macromolecular complexation between bovine serum albumin and the self-assembled hydrogel nanoparticle of hydrophobized polysaccharides, J Am Chem Soc, 1996, 118(26), 6110–15. [183] Ouchi T, Nishizawa H, Ohya Y. Aggregation phenomenon of PEG-grafted chitosan in aqueous solution, Polymer, 1998, 39(21), 5171–75.

References

| 187

[184] Yoksan R, Matsusaki M, Akashi M, Chirachanchai S. Controlled hydrophobic/hydrophilic chitosan: colloidal phenomena and nanosphere formation, Colloid Polym Sci, 2004, 282(4), 337–42. [185] Choisnard L, Geze A, Putaux J, Wong Y, Wouessidjewe D. Nanoparticles of β-cyclodextrin esters obtained by self-assembling of biotransesterified β-cyclodextrins, Biomacromolecules, 2006, 7(2), 515–20. [186] Wang Y, Liu L, Jiang Q, Zhang Q. Self-aggregated nanoparticles of cholesterol-modified chitosan conjugate as a novel carrier of epirubicin, Eur Polym J, 2007, 43(1), 43–51. [187] Park K, Kim J, Nam YS, Lee S, Nam HY, Kim K, Park JH, Kim I, Choi K, Kim SY, Others. Effect of polymer molecular weight on the tumor targeting characteristics of self-assembled glycol chitosan nanoparticles, J Control Release, 2007, 122(3), 305–14. [188] Bravo-Osuna I, Millotti G, Vauthier C, Ponchel G. In vitro evaluation of calcium binding capacity of chitosan and thiolated chitosan poly (isobutyl cyanoacrylate) core-shell nanoparticles, Int J Pharmaceut, 2007, 338(1), 284–90. [189] Bravo-Osuna I, Ponchel G, Vauthier C. Tuning of shell and core characteristics of chitosandecorated acrylic nanoparticles, Eur J Pharm Sci, 2007, 30(2), 143–54. [190] Vila A, Sánchez A, Tob O M, Calvo P, Alonso MJ. Design of biodegradable particles for protein delivery, J Control Release, 2002, 78(1–3), 15–24. [191] Singnurkar PS, Gidwani SK. Evaluation of hydrophobic nanoparticulate delivery system for insulin, Indian J Pharm Sci, 2008, 70(6), 721. [192] Reis CP, Ribeiro AONJ, Houng S, Veiga F, Neufeld RJ. Nanoparticulate delivery system for insulin: design, characterization and in vitro/in vivo bioactivity, Eur J Pharm Sci, 2007, 30(5), 392–97. [193] Cetin M, Aktas Y, Vural I, Capan Y, Dogan LA, Duman M, Dalkara T. Preparation and in vitro evaluation of bfgf-loaded chitosan nanoparticles, Drug Deliv, 2007, 14(8), 525–29. [194] Li Y, Nagira T, Tsuchiya T. The effect of hyaluronic acid on insulin secretion in Hit-T15 cells through the enhancement of gap-junctional intercellular communications, Biomaterials, 2006, 27(8), 1437–43. [195] Azizi E, Namazi A, Haririan I, Fouladdel S, Khoshayand MR, Shotorbani PY, Nomani A, Gazori T. Release profile and stability evaluation of optimized chitosan/alginate nanoparticles as egfr antisense vector, Int J Nanomed, 2010, 5, 455. [196] Elzoghby AO, Samy WM, Elgindy NA. Albumin-based nanoparticles as potential controlled release drug delivery systems, J Control Release, 2012, 157(2), 168–82. [197] Reis CP, Ribeiro AONJ, Veiga F, Neufeld RJ, Damg E C. Polyelectrolyte biomaterial interactions provide nanoparticulate carrier for oral insulin delivery, Drug Deliv, 2008, 15(2), 127–39. [198] Sarmento B, Martins S, Ribeiro AON, Veiga F, Neufeld R, Ferreira D. Development and comparison of different nanoparticulate polyelectrolyte complexes as insulin carriers, Int J Pept Res Ther, 2006, 12(2), 131–38. [199] Pan Y, Li Y, Zhao H, Zheng J, Xu H, Wei G, Hao J, Others. Bioadhesive polysaccharide in protein delivery system: chitosan nanoparticles improve the intestinal absorption of insulin in vivo, Int J Pharmaceut, 2002, 249(1), 139–47. [200] Myrick JM, Vendra VK, Krishnan S. Self-assembled polysaccharide nanostructures for controlled-release applications, Nanotechnology Reviews, 2014, f3(4), 319–46. [201] Na K, Bum Lee T, Park K, Shin E, Lee Y, Choi H. Self-assembled nanoparticles of hydrophobically-modified polysaccharide bearing vitamin h as a targeted anti-cancer drug delivery system, Eur J Pharm Sci, 2003, 18(2), 165–73. [202] Sarmah JK, Bhattacharjee SK, Mahanta R, Mahanta R. Preparation of cross-linked guar gum nanospheres containing tamoxifen citrate by single step emulsion in situ polymer crosslinking method, J Incl Phenom Macro, 2009, 65(3–4), 329–34.

188 | 8 Novel nanoparticle materials for drug/food delivery-polysaccharides

[203] Lu B, Xiong S, Yang H, Yin X, Zhao R. Mitoxantrone-loaded BSA nanospheres and chitosan nanospheres for local injection against breast cancer and its lymph node metastases: I: formulation and in vitro characterization, Int J Pharmaceut, 2006, 307(2), 168–74. [204] Yang X, Zhang Q, Wang Y, Chen H, Zhang H, Gao F, Liu L. Self-aggregated nanoparticles from methoxy poly(ethylene glycol)-modified chitosan: synthesis; characterization; aggregation and methotrexate release in vitro, Colloids and Surfaces B: Biointerfaces, 2008, 61(2), 125–31. [205] Zhang J, Chen XG, Li YY, Liu CS. Self-assembled nanoparticles based on hydrophobically modified chitosan as carriers for doxorubicin, Nanomedicine: Nanotechnology, Biology and Medicine, 2007, 3(4), 258–65. [206] Hu F, Ren G, Yuan H, Du Y, Zeng S. Shell cross-linked stearic acid grafted chitosan oligosaccharide self-aggregated micelles for controlled release of paclitaxel, Colloids and Surfaces B: Biointerfaces, 2006, 50(2), 97–103. [207] Di Gioia S, Trapani A, Castellani S, Carbone A, Belgiovine G, Craparo EF, Puglisi G, Cavallaro G, Trapani G, Conese M. Nanocomplexes for gene therapy of respiratory diseases: targeting and overcoming the mucus barrier, Pulm Pharmacol Ther, 2015, 34, 8–24. [208] Mansouri S, Lavigne P, Corsi K, Benderdour M, Beaumont E, Fernandes JC. Chitosan-DNA nanoparticles as non-viral vectors in gene therapy: strategies to improve transfection efficacy, Eur J Pharm Biopharm, 2004, 57(1), 1–08. [209] Katas H, Alpar HO. Development and characterisation of chitosan nanoparticles for sirna delivery, J Control Release, 2006, 115(2), 216–25. [210] Kaul G, Amiji M. Long-circulating poly(ethylene glycol)-modified gelatin nanoparticles for intracellular delivery, Pharm Res, 2002, 19(7), 1061–67. [211] Liu Q, Wang C, Cao Y, Xu X, Zhang L. A novel gene carrier prepared from triple helical β-glucan and polydeoxyadenylic acid, J Mater Chem B, 2014, 2(8), 933–44. [212] Jiménez-Colmenero F. Potential applications of multiple emulsions in the development of healthy and functional foods, Food Res Int, 2013, 52(1), 64–74. [213] Jeddi-Tehrani M, Mahmoudi AR, Sabzvari A, Atyabi F, Dinarvand R. Stabilization of monoclonal antibody upon encapsulation in polymeric nanoparticles by double emulsion technique, J Control Release, 2013, 172(1), e62–63. [214] Trombino S, Cassano R, Muzzalupo R, Pingitore A, Cione E, Picci N. Stearyl Ferulate-based solid lipid nanoparticles for the encapsulation and stabilization of β-carotene and αtocopherol, Colloids and Surfaces B: Biointerfaces, 2009, 72(2), 181–87. [215] Koudelka S, Turanek Knotigova P, Masek J, Prochazka L, Lukac R, Miller AD, Neuzil J, Turanek J. Liposomal delivery systems for anti-cancer analogues of vitamin E, J Control Release, 2015, 207, 59–69. [216] Polyakov NE, Kispert LD. Water soluble biocompatible vesicles based on polysaccharides and oligosaccharides inclusion complexes for carotenoid delivery, Carbohyd Polym, 2015, 128, 207–19. [217] Wicki A, Witzigmann D, Balasubramanian V, Huwyler JR. Nanomedicine in cancer therapy: challenges, opportunities, and clinical applications, J Control Release, 2015, 200, 138–57. [218] Voordouw J, Antonides G, Cornelisse-Vermaat JR, Pfaff S, Niemietz D, Frewer LJ. Optimising the delivery of food allergy information. an assessment of food allergic consumer preferences for different information delivery formats, Food Qual Prefer, 2012, 23(1), 71–78. [219] Liu X, Liu C, Zhang W, Xie C, Wei G, Lu W. Oligoarginine-modified biodegradable nanoparticles improve the intestinal absorption of insulin, Int J Pharmaceut, 2013, 448(1), 159–67. [220] Gaumet M, Gurny R, Delie F. Interaction of biodegradable nanoparticles with intestinal cells: the effect of surface hydrophilicity, Int J Pharmaceut, 2010, 390(1), 45–52.

References

|

189

[221] Hu B, Ting Y, Zeng X, Huang Q. Cellular uptake and cytotoxicity of chitosan– caseinophosphopeptides nanocomplexes loaded with epigallocatechin gallate, Carbohyd Polym, 2012, 89(2), 362–70. [222] Xiao J, Nian S, Huang Q. Assembly of kafirin/carboxymethyl chitosan nanoparticles to enhance the cellular uptake of curcumin, Food Hydrocolloid, 2015, 51, 166–75. [223] Anand P, Nair HB, Sung B, Kunnumakkara AB, Yadav VR, Tekmal RR, Aggarwal BB. Design of curcumin-loaded plga nanoparticles formulation with enhanced cellular uptake, and increased bioactivity in vitro and superior bioavailability in vivo, Biochem Pharmacol, 2010, 79(3), 330–38. [224] Amidi M, Mastrobattista E, Jiskoot W, Hennink WE. Chitosan-based delivery systems for protein therapeutics and antigens, Adv Drug Deliver Rev, 2010, 62(1), 59–82. [225] Rhim J, Park H, Ha C. Bio-nanocomposites for food packaging applications, Prog Polym Sci, 2013, 38(10–11), 1629–52. [226] Sorrentino A, Gorrasi G, Vittoria V. Potential perspectives of bio-nanocomposites for food packaging applications, Trends Food Sci Tech, 2007, 18(2), 84–95. [227] Kumari A, Yadav SK, Pakade YB, Singh B, Yadav SC. Development of biodegradable nanoparticles for delivery of quercetin, Colloids and Surfaces B: Biointerfaces, 2010, 80(2), 184–92. [228] Matalanis A, Decker EA, McClements DJ. Inhibition of lipid oxidation by encapsulation of emulsion droplets within hydrogel microspheres, Food Chem, 2012, 132(2), 766–72. [229] Matalanis A, McClements DJ. Hydrogel microspheres for encapsulation of lipophilic components: optimization of fabrication & performance, Food Hydrocolloid, 2013, 31(1), 15–25. [230] Chiu C, Lin J. Self-assembly behavior of polymer-assisted clays, Prog Polym Sci, 2012, 37(3), 406–44. [231] Mun S, Kim Y, Shin M, McClements DJ. Control of lipid digestion and nutraceutical bioaccessibility using starch-based filled hydrogels: influence of starch and surfactant type, Food Hydrocolloid, 2015, 44, 380–89. [232] McClements DJ, Li Y. Structured emulsion-based delivery systems: controlling the digestion and release of lipophilic food components, Adv Colloid Interfac, 2010, 159(2), 213–28. [233] Matos M, Timgren A, Sj M, Dejmek P, Rayner M. Preparation and encapsulation properties of double pickering emulsions stabilized by quinoa starch granules, Colloids and Surfaces A: Physicochemical and Engineering Aspects, 2013, 423, 147–53. [234] Sapei L, Naqvi MA, Rousseau D. Stability and release properties of double emulsions for food applications, Food Hydrocolloid, 2012, 27(2), 316–23. [235] Peres I, Rocha S, Gomes J, Morais S, Pereira MC, Coelho M. Preservation of catechin antioxidant properties loaded in carbohydrate nanoparticles, Carbohyd Polym, 2011, 86(1), 147–53.

Index Acinebacter sp. ADP1 51 alkanes 51 applicability 50 applications 89

genome 47 genomic sequence 46 genomics 46 global problems 45

bacterial DyP 108 bagasse 18, 30, 38 biobutanol 52 biodiesel 16, 21, 23, 50 biofuel 16–18, 29, 30, 45, 50 biomass 45 biopharmaceutical 31 biosynthetic pathway 81 biotechnological applications 122 butanol 53

heme protein 107 high-throughput techniques 48

13 C-labelled

49 catalytic property 108 cell factories 45, 53 cell-wide 45 cellular metabolism 46 chondroitin 23 chromatography 1, 8, 10–12 Clostridium acetobutylicum 52 computational methods 49 conclusion 53 cyanobacteria 51 dimer 117, 118 dimerization 117 distal heme residue 111 dye decolorization 123 dye-decolorizing peroxidases 107 E. coli 50, 52 economics 1, 2, 8, 10, 11, 25, 30, 33, 35, 38 electricity 1, 17, 18, 20, 21 engineered 32 ethanol 6, 16, 20, 23, 27, 29, 30 factors 53 FAEE 50 Farnesane 51 fluxome 49 fluxomics 49

in silico simulation 50 jet fuel 51 lignin 2, 17, 28, 35 lignin degradation 111 lignin valorization 122 ligno-cellulosic 17–19, 25–30, 35 manganese peroxidase 113 mathematical modeling 49 metabolic engineering 45, 53, 88 metabolic engineering strategies 50 metabolic network 53 metabolic pathways 52 metabolites 48 metabolomics 48 mRNA 48 nanofiber 35–38 olefins 51, 52 OleTJE 52 omics 46 outlook 53 paper 5, 17–20, 28, 36, 37 peroxidase catalytic cycle 114 pharmaceutical 1–3, 6, 8, 11, 14, 15 physiological roles 125 production process 76 protein 48 protein expression 47 proteomics 48 proximal histidine 118 pulp 5, 17, 18, 28, 36 reaction intermediates 114

192 | Index

S. cerevisiae 51 sphingans 73 structure-function relationship 117 substrate specificity 108 synthetic biology 45 system-level modeling 46 systems biology 46, 53

transcription 47 transcriptomics 47 transient kinetics 119 welan gum 75 Zymomonas mobilis 50