Biocatalysis and Biomolecular Engineering E-Book

• Herausgeber: John Wiley & Sons
• Kategorie: Wissenschaft und neue Technologien
• Sprache: Englisch

Seitenzahl: 801

Veröffentlichungsjahr: 2010

Beschreibung

An expert overview of new technologies guiding the construction of a sustainable society

This compendium of important insights from sixty distinguished international scholars looks at the significant advances in progressive environmental technology—especially the molecular engineering used on plants, animals, and microorganisms—as the game changer in the high-stakes race to reverse earth-damaging practices.

Biocatalysis and Biomolecular Engineering covers subject matter on the latest developments in eco-friendly and energy-saving manufacturing processes with the emphasis on agricultural technology and bio-based products. Focusing its study on remedies that show promise in curing food and energy ills, this book examines groundbreaking work in various fields, such as nutraceuticals, genetic engineering of agricultural products, and bioenergy. Biocatalysis and Biomolecular Engineering:

• Can be used as a reference by teachers, graduate students, and industrial scientists who conduct research in bioscience and biotechnology

• Serves as the first book to bring together fundamentals and leading-edge technologies for the development of bio-based industrial products through biocatalysis; for example, it discusses the preparation of biofunctional micro- and nanoparticles

• Contains chapters by international experts from academia, industry, and government research institutes

Biocatalysis and Biomolecular Engineering builds a cohesive, well thought out case for nurturing new discoveries in eco-technology by inviting critical discussion on devising viable solutions to sustaining the future wellness of humankind.

Leseprobe

Copyright

Copyright © 2010 by John Wiley & Sons, Inc. All rights reserved.

Published by John Wiley & Sons, Inc., Hoboken, New Jersey

Published simultaneously in Canada

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Library of Congress Cataloging-in-Publication Data:

ISBN: 9780470487594

To our wives, Mandy Hou and Yea-Shiow Shaw, for their understanding and kind support during preparation of this book

Contents

Cover

Title Page

Copyright

Preface

Contributors

Section I Improvement of Agronomic and Microbial Traits

1 I Insights into the Structure and Function of Acyl-CoA: Diacylglycerol Acyltransferase

1.1 Introduction

1.2 Discovery of DGAT

1.3 Membrane Topological Organization of DGATs

1.4 Alignment of DGAT1 Polypeptides

1.5 Alignment of DGAT2 Polypeptides

1.6 Structure of DGAT Genes

1.7 Functional Motifs in DGAT1

1.8 Functional Motifs in DGAT2

1.9 Subcellular Localization of DGATs

1.10 Conclusions and Future Research

Acknowledgments

Abbreviations

References

2 Improving Enzyme Character by Molecular Breeding: Preparation of Chimeric Genes

2.1 Introduction

2.2 Preparation of Chimeric β-Glucosidase to Improve the Enzyme Character

2.3 Preparation of Chimeric Xylanase to Determine the Enzyme Activity at Basic pH

2.4 Future Studies

References

3 Production and Accumulation of Unusual Fatty Acids in Plant Tissues

3.1 Introduction

3.2 Results and Discussion

3.3 Conclusions

References

4 Preparation of Oleaginous Yeast by Genetic Modification and Its Potential Applications

4.1 Introduction

4.2 Identification of Genes Involved in Lipid Accumulation in S. cerevisiae

4.3 Preparation of Oleaginous Yeast S. cerevisiae by Genetic Modification

4.4 Future Perspectives

References

5 Improving Value of Oil Palm Using Genetic Engineering

5.1 Introduction

5.2 Materials and Methods

5.3 Results and Discussion

Acknowledgments

References

6 Potential in Using Arabidopsis Acyl-Coenzyme-A-Binding Proteins in Engineering Stress-Tolerant Plants

6.1 Plant Lipid-Binding Proteins

6.2 Proposed Biological Roles of Plant LTPs

6.3 Arabidopsis ACBPs

6.4 Potential of Arabidopsis ACBP1 in Phytoremediation

6.5 Potential of Arabidopsis ACBP6 in Enhancing FREEZING Tolerance

6.6 Potential of Arabidopsis ACBP2 in Combating Oxidative Stress

6.7 Conclusions and Perspectives

Acknowledgments

References

7 Modification of Lipid Composition by Genetic Engineering in Oleaginous Marine Microorganism, Thraustochytrid

7.1 Introduction

7.2 Materials and Methods

7.3 Results and Discussion

References

8 Integrated Approaches to Manage Tomato Yellow Leaf Curl Viruses

8.1 Introduction

8.2 Host-Plant Resistance to TYLCV

8.3 Resistance to the Whitefly Vector

8.4 Pathogen-Derived Resistance

8.5 Integrated Approach Towards Stable Resistance to TYLCV

8.6 Conclusions

References

9 Carbohydrate Acquisition During Legume Seed Development

9.1 Introduction: Legume Seed Crops

9.2 Nutrient Pathway from Seed Coat to Embryo in Legumes

9.3 Role of Invertases in Sucrose Metabolism During Legume Seed Development

9.4 Pea Seed Coat Morphology And Site of Sucrose Unloading

9.5 Embryo Acquisition of Sugars

9.6 Modification of Nutrient Pathway During Pea Seed Development

References

10 Biotechnology Enhancement of Phytosterol Biosynthesis in Seed Oils

10.1 Introduction

10.2 Occurrence and Levels of Phytosterol in Seed Oils

10.3 Phytosterol Enhancement Through Gene Engineering

10.4 Remarks and Perspectives

References

Section II Functional Foods and Biofuels

11 Dietary Phosphatidylinositol in Metabolic Syndrome

11.1 Introduction

11.2 Effect of Dietary PI on the Development of Non-alcoholic Fatty Liver Disease in Metabolic Syndrome Model Rats

11.3 Effect of Dietary PI on Serum Adiponectin Levels and Hepatic Inflammatory Molecule mRNA Levels in Metabolic Syndrome Model Rats

11.4 Effect of Dietary PI on Cholesterol Levels in Metabolic Syndrome Model Rats

11.5 Effect of Dietary PI on Hepatic mRNA Levels and Fecal Bile Acid Levels in Metabolic Syndrome Model Rats

11.6 Conclusions

Acknowledgment

References

12 Biotechnological Enrichment of Cereals with Polyunsaturated Fatty Acids

12.1 Introduction

12.2 Importance and Sources of Polyunsaturated Fatty Acids

12.3 Biotechnological Strategy for Cereals Enriched with PUFAs

12.4 Solid-State Fermentations

12.5 Genetic Transformation of Plants

12.6 Conclusions and Perspectives

Acknowledgment

References

13 Lipophilic Ginsenoside Derivatives Production

13.1 Introduction

13.2 Chemical Composition and Traditional Usage of Panax ginseng

13.3 Metabolism of Ginsenosides in Human Body

13.4 Production of Lipophilic Ginsenoside Derivatives Using Enzyme

References

14 Brown Seaweed Lipids as Possible Source for Nutraceuticals and Functional Foods

14.1 Introduction

14.2 Seaweeds Lipids as a Rich Source of Functional HUFA

14.3 Brown Seaweeds are a Rich Source of Polyphenols

14.4 Carotenoids in Brown Seaweeds

14.5 Future Direction

References

15 Processes for Production of Biodiesel Fuel

15.1 Introduction

15.2 Biofuels

15.3 Various Processes for BDF Production

15.4 Processes with Chemical Catalysts

15.5 Processes without Catalysts

15.6 Biochemical Processes for BDF Production

15.7 Conclusions

References

16 Noncatalytic Alcoholysis Process for Production of Biodiesel Fuel: Its Potential in Japan and Southeast Asia

16.1 Promising Materials for the Production of Biodiesel Fuel

16.2 Problems with the Conventional Alkaline Catalyzed Alcoholysis Reaction Process for the Production of Biodiesel Fuel

16.3 Advantages of a Noncatalytic Alcoholysis Reaction Process Over the Conventional Alkaline-Catalyzed Process

16.4 Research on the Noncatalytic Alcoholysis Reaction for Biodiesel Fuel Production in Japan

16.5 Conclusions

Acknowledgment

References

17 Use of Coniochaeta ligniaria to Detoxify Fermentation Inhibitors Present in Cellulosic Sugar Streams

17.1 Introduction

17.2 Discovery of Microorganisms for Abatement of Fermentation Inhibitors

17.3 C. ligniaria Metabolism of Inhibitors

17.4 Bioabatement of Pretreated Hydrolysates

17.5 Conclusions

References

18 Omics Applications to Biofuel Research

18.1 Introduction

18.2 Next Generation: Renewable Energy Biomass Program

18.3 Main Feedstocks for Next-Generation Biofuels

18.4 Identification of Cellulase Genes by Genomic Approaches

18.5 How Can Transcriptomic Study Help Identify Cellulase Genes in a Microbe?

18.6 How Can Proteomic Study Help Identify Cellulase Genes in a Microbe?

18.7 Conversion of Cellulose to Ethanol

18.8 Concluding Remarks

References

Section III Renewable Bioproducts

19 Biotechnological Uses of Phospholipids

19.1 Introduction

19.2 Phospholipids for Food and Nutraceuticals

19.3 Phospholipids for Cosmetics

19.4 Phospholipids for Agricultural Application

19.5 Phospholipids for Pharmaceuticals

19.6 Conclusions

References

20 Application of Partition Chromatographic Theory on the Routine Analysis of Lipid Molecular Species

20.1 Relationship between the Structure of a Lipid Molecule and the Sequence of Elution: The Traditional Way of Predicting the Retention Time on Reverse-Phase HPLC

20.2 Relationship between the Structure of a Lipid Molecule and the Sequence of Elution: Predicting the Retention Time of Individual Lipid Molecular Species on Reverse-Phase HPLC

20.3 Application of the Simple Additional Theorem of Chemical Potential in the Chromatographic System

20.4 Calculation of Relative Retention Potential Index (RPI) on HPLC

Appendixes

Acknowledgments

References

21 Dehydrogenase-Catalyzed Synthesis of Chiral Intermediates for Drugs

21.1 Introduction

21.2 Dehydrogenase-Catalyzed Reductions

21.3 Dehydrogenase-Catalyzed Reductive Aminations

21.4 Conclusions

References

22 Engineering of Bacterial Cytochrome P450 Monooxygenase as Biocatalysts for Chemical Synthesis and Environmental Bioremediation

22.1 Introduction

22.2 H2O2-Dependent Substrate Hydroxylation Activity and H2O2 Inactivation of Mutant Cytochrome P450 BM-3

22.3 Engineering Cytochrome P450 BM-3 for Oxidation of Polycyclic Aromatic Hydrocarbons

22.4 Metabolism of Polychlorinated Dibenzo-p-Dioxins by Cytochrome P450 BM-3 and its Mutant

22.5 Stereoselectivity in Propylbenzene and 3-Chlorostyrene Oxidation by Cytochrome P450 BM-3 and its Mutant

22.6 Application of Cytochrome P450 BM-3 Mutant to the Synthesis of Hydroquinone Derivatives from Phenolic Compounds

22.7 Conclusions

References

23 Glycosynthases from Inverting Hydrolases

23.1 Retaining and Inverting Glycoside Hydrolases

23.2 Reactions of GH with the Glycosyl Fluoride of the Opposite Anomer

23.3 The Model Inverting GH: Reducing-End-Xylose Releasing Exo-Oligoxylanase (Rex)

23.4 Hehre Resynthesis-Hydrolysis of Rex

23.5 Conversion of Rex into Glycosnythase by Mutating Base Residue

23.6 Conversion of Rex into Glycosnythase by Mutating the Residue Supporting the Nucleophilic Water Molecule

23.7 Comparison of Glycosynthase Conversion from Retaining and Inverting GHs

23.8 Glycosynthase from an Inverting GH with A typical Reaction Mechanism

References

24 Molecular Species of Diacylglycerols and Triacylglycerols Containing Dihydroxy Fatty Acids in Castor Oil

24.1 Introduction

24.2 HPLC Fractionation of the Molecular Species of Acylglycerols in Castor Oil

24.3 Proposed Structures of Dihydroxy Fatty Acids

24.4 Structures of Acylglycerols Containing Dihydroxy Fatty Acids

24.5 Regiospecific Quantification of Triacylglycerols

24.6 Ratios of Fatty Acids at the SN-2 Position of Triacylglycerols

24.7 Conclusions

References

25 Biocatalytic Production of Lactobionic Acid

25.1 Introduction

25.2 Practical and Feasible Applications

25.3 Biocatalytic Production Methods

25.4 Production by Paraconiothyrium Oxidase

25.5 Conclusions

References

26 Recent Advances in Aldolase-Catalyzed Synthesis of Unnatural Sugars and Iminocyclitols

26.1 Introduction

26.2 Directed Evolution of L-Rhamnulose 1-Phosphate Aldolase Using In Vivo Selection

26.3 Use of Borate as a Phosphate Ester Mimic in Aldolase-Catalyzed Reactions: Practical Synthesis of L-Fructose and L-Iminocyclitols

26.4 One-Pot Synthesis of D-Iminocyclitols Using D-Fructose 6-Phosphate Aldolase

26.5 Conclusions

Acknowledgments

References

27 Production of Value-Added Products by Lactic Acid Bacteria

27.1 Introduction

27.2 Lactate Fermentation

27.3 Production of Antibacterial Peptides and Proteins

27.4 Other Applications

27.5 Perspectives

References

28 Enzymatic Synthesis of Glycosides Using Alpha-Amylase Family Enzymes

28.1 Introduction

28.2 Enzymatic Synthesis of Alpha-Arbutin and its Melanogenesis Inhibition

28.3 Enzymatic Syntheses of Hydroquinone Glycosides and their Inhibitory Effects on Human Tyrosinase

28.4 Transglycosylation to Carboxylic Acid by Sucrose Phosphorylase

28.5 Conclusions

References

29 Biological Synthesis of Gold and Silver Nanoparticles Using Plant Leaf Extracts and Antimicrobial Application

29.1 Introduction

29.2 Synthesis of Silver Nanoparticles Using Plant Leaf Extracts

29.3 Synthesis of Gold Nanoparticles Using Plant Leaf Extracts

29.4 Synthesis of Gold/Silver Bimetal Nanoparticles Using Plant Leaf Extracts

29.5 Antimicrobial Application of Silver Nanoparticles Synthesized Using Plant Leaf Extracts

29.6 Conclusions

References

30 Potential Approach of Microbial Conversion to Develop New Antifungal Products of Omega-3 Fatty Acids

30.1 Introduction

30.2 Development of New Antifungal Products of Omega-3 Fatty Acid by the Microbial Conversion

30.3 Biological Activities of Microbially Converted New Antifungal Products of Omega-3 Fatty Acids

30.4 Conclusions

Acknowledgment

References

Index

Preface

This book was assembled with the intent of bringing together current advances and in-depth reviews of biocatalysis and biomolecular engineering with emphasis on agricultural biotechnology. The book consists of selected papers presented at the fourth International Symposium on Biocatalysis and Biotechnology held at the Academia Sinica, Taipei, Taiwan November 19–21, 2008. At this symposium, 60 distinguished international scientists from the United States, Japan, Korea, Canada, Brazil, Belgium, Slovak Republic, France, and Taiwan, shared their valuable research results. Additionally, there were 20 selected posters, one session for American Oil Chemists Society Asian Section, and two workshops for Biotech Developments and over 600 attendees. A few chapters contained in this book were contributed by distinguished scientists who could not attend this meeting. This meeting was a great success and we greatly appreciate President Dr. Chi-Huey Wong of Academia Sinica for providing the venue for the meeting. The contributions of local organization committee members are highly appreciated: Andrew H.-J. Wang, and Ming-Che Shih of Academia Sinica, and Yung-Sheng Huang, Chang-Hsien Yang of the National Chung Hsing University.

Recent energy and food crises point out the important of bio-based products from renewable resources and agricultural biotechnology. It is inevitable to use modern tools of molecular engineering on plants, animals and microorganisms to solve these crises and improve the wellness of humankind. There is no comprehensive book on molecular engineering of agricultural biotechnology and bio-based products from renewable resources. The authors are internationally recognized experts from all sectors of academia, industry, and government research institutes. This is the most current book on molecular engineering of agricultural biotechnology and bio-based industrial products.

This book composes of 30 chapters divided into three sections. The first 10 chapters describe the world's newest research on improvement of agronomic and microbial traits. Included are: Insights into the Structure and Function of Acyl-CoA: Diacylglycerol Acyltransferase, Improving Enzyme Character by Molecular Breeding-Preparation of Chimeric Genes, Production and Accumulation of Unusual Fatty Acids in Plant Tissues, Preparation of Oleaginous Yeast by Genetic Modification and Its Potential Applications, Improving Value of Oil Palm Using Genetic Engineering, Potential in Using Arabidopsis Acyl-Coenzyme-A-Binding Proteins in Engineering Stress-Tolerant Plants, Modification of Lipid Composition by Genetic Engineering in Oleaginous Marine Microorganisms: Thraustochytrid, Integrated Approaches to Manage Tomato Yellow Leaf Curl Viruses, Carbohydrate Acquisition During Legume Seed Development, and Biotechnology Enhancement of Phytosterol Biosynthesis in Seed Oils. The second section includes 8 chapters devoted to Functional Foods and Biofuels: Dietary Phosphatydyl Inositol in Metabolic Syndrome, Biotechnological Enrichment of Cereals with Polyunsaturated Fatty Acids, Brown Seaweeds Lipids as Possible Source for Nutraceuticals and Functional Foods, Lipophillic Ginsenosides Derivative Production, Processes for Production of Biodiesel Fuel, Noncatalytic Alcoholysis Process for Production of Biodiesel Fuel—Its Potential in Japan and Southeast Asia, Use of Coniochaeta Ligniaria to Detoxify Fermentation Inhibitors Present in Cellulosic Sugar Streams, and Omics Applications to Biofuel Research. The third section with 12 chapters describes Renewable Bioproducts: Biotechnological Uses of Phospholipids, Application of Partition Chromatographic Theory on the Routine Analysis of Lipid Molecular Species, Dehydrogenase-Catalyzed Synthesis of Chiral Intermediates for Drugs, Engineering of Bacterial Cytochrome P450 Monooxygenase as Biocatalysts for Chemical Synthesis and Environmental Bioremediation, Glycosynthase from Inverting Hydrolases, Molecular Species of Diacylglycerols and Triacylglycerols Containing Dihydroxy Fatty Acids in Castor Oil, Biocatalytic Production of Lactobionic Acid, Recent Advances in Aldolase-Catalyzed Synthesis of Unnatural Sugars and Iminocyclitols, Production of Value-Added Products by Lactic Acid Bacteria, Enzyme Synthesis of Glycosides Using Alpha-Amylase Family Enzymes, Biological Synthesis of Gold and Silver Nanoparticles Using Plant Leaf Extracts and Antimicrobial Application, and Potential Approach of Microbial Conversion to Develop New Antifungal Products of Omega-3 Fatty Acids.

This book serves as reference for teachers, graduate students, and industrial scientists who conduct research in biosciences and biotechnology.

Ching T. Hou

Peoria, IL USA

Jei-Fu Shaw

Taichung, Taiwan

Contributors

Hirofumi Adachi, Department of Molecular Biotechnology, Graduate School of Advanced Sciences of Matter, Hiroshima University, 1-3-1 Kagamiyama, Higashi-Hiroshima, Japan

Zuzana Adamechova, Department of Biochemical Technology, Faculty of Chemical and Food Technology, Slovak University of Technology, Radlinskeho 9, Bratislava, Slovak Republic

Tsunehiro Aki, Department of Molecular Biotechnology, Hiroshima University, 1-3-1 Kagamiyama, Higashi-Hiroshima, Japan

Nur Hanin Ayub, Malaysian Palm Oil Board, No. 6, Persiaran Institusi, Bandar Baru Bangi, Kajang, Selangor, Malaysia

Hassan Azaizeh, Institute of Applied Research Center (Affiliated with University of Haifa), The Galilee Society, Shefa-Amr, Israel

Vivek K. Bajpai, Department of Biotechnology, Daegu University, Kyoungsan, Kyoungbook, Republic of Korea

Pankaj K. Bhowmik, National Research Council—Plant Biology Institute Saskatoon, Saskatchewan, Canada

Kenneth M. Bischoff, Renewable Product Technology Research Unit, National Center for Agricultural Utilization Research, Agricultural Research Service, United States Department of Agriculture, 1815 N. University St., Peoria, IL, USA

Bahariah Bohari, Malaysian Palm Oil Board, No. 6, Persiaran Institusi, Bandar Baru Bangi, Kajang, Selangor, Malaysia

Milan Certik, Department of Biochemical Technology, Faculty of Chemical and Food Technology, Slovak University of Technology, Bratislava, Slovak Republic

Hsin-Liang Chen, Biodiversity Research Center, Academia Sinica, Taipei, Taiwan

Qin-Fang Chen, School of Biological Sciences, The University of Hong Kong, Pokfulam Road, Hong Kong, China

Qilin Chen, National Research Council Canada—Plant Biotechnology Institute, 110 Gymnasium Place, Saskatoon, Saskatchewan, Canada

Mee-Len Chye, School of Biological Sciences, The University of Hong Kong, Pokfulam, Hong Kong and State (China) Key Laboratory of Agrobiotechnology, Chinese University of Hong Kong, Shatin, Hong Kong

Fengjie Cui, Department of Food, Agricultural, and Biological Engineering, The Ohio State University/Ohio Agricultural Research and Development Center, Wooster, USA

Robert de la Pena, AVRDC—The World Vegetable Center, Shanhua, Tainan, Taiwan

Bruce S. Dien, Bioenergy Research Unit, NCAUR, USDA-ARS, Peoria, IL, 61604, USA

Ahmad Kushairi Din, Malaysian Palm Oil Board, No. 6, Persiaran Institusi, Bandar Baru Bangi, Kajang, Selangor, Malaysia

Wei Gao, School of Biological Sciences, The University of Hong Kong, Pokfulam Road, Hong Kong, China

William A. Greenberg, Department of Chemistry, The Scripps Research Institute, 10550, N. Torrey Pines Rd., La Jolla, CA 92037, USA

Shoji Hagiwara, National Food Research Institute, NARO, Kan-nondai, Tsukuba, Ibaraki, Japan

J. J. Han, Doosan Glonet, B5F, Advanced Convergence Institute of Technology, 864-1 IUI-dong, Suwon, Gyeonggi, Korea

Ahmad Tarmizi Hashim, Malaysian Palm Oil Board, No. 6, Persiaran Institusi, Bandar Baru Bangi, Kajang, Selangor, Malaysia

T. Hatanaka, Kobe University, Kobe, Japan

Kiyoshi Hayashi, National Food Research Institute, 2-1-12 Kan-nondai, Tsukuba, Ibaraki, Japan

Xiaohua He, Western Regional Research Center, United States Department of Agriculture, Albany, California, USA

David Hildebrand, Agronomy Department, University of Kentucky, Lexington, Kentucky, USA

Tsugihiko Hirano, Renesas Northern Japan Semiconductor, Inc., 145-1 Nakajima Nanae-cho Kameda-gun, Hokkaido, Japan

Zhangyong Hong, Department of Chemistry, The Scripps Research Institute, 10550, N. Torrey Pines Rd., La Jolla, CA 92037, USA

Masashi Hosokawa, Faculty of Fisheries Sciences, Hokkaido University, 3-1-1 Minato, Hakodate, Japan

Ching T. Hou, Microbial Genomic and Bioprocessing Research Unit, National Centre for Agricultural Utilization Research, ARS, USDA, Peoria, IL, USA

Jiang-Ning Hu, Department of Food Science and Technology, Chungnam National University, 220 Yusung Gu, Gung-Dong, Daejeon, Republic of Korea

J. Hughes, AVRDC—The World Vegetable Center, Shanhua, Tainan, Taiwan

Zamzuri Ishak, Malaysian Palm Oil Board, No. 6, Persiaran Institusi, Bandar Baru Bangi, Kajang, Selangor, Malaysia

Hiroaki Iwasaka, Department of Molecular Biotechnology, Graduate School of Advanced Sciences of Matter, Hiroshima University, 1-3-1 Kagamiyama, Higashi-Hiroshima, Japan

Ghulam Kadir Ahmad Parveez, Malaysian Palm Oil Board, No. 6, Persiaran Institusi, Bandar Baru Bangi, Kajang, Selangor, Malaysia

P. Kadirvel, AVRDC—The World Vegetable Center, Shanhua, Tainan 71499, Taiwan

Toshihide Kakizono, Department of Molecular Biotechnology, Graduate School of Advanced Sciences of Matter, Hiroshima University, 1-3-1 Kagamiyama, Higashi-Hiroshima, Japan

Yasushi Kamisaka, Institute for Biological Resources and Functions, National Institute of Advanced Industrial Science and Technology, Japan

Sun Chul Kang, Department of Biotechnology, College of Engineering, Daegu University, Gyungsan City, Gyungbook, Korea

Seiji Kawamoto, Department of Molecular Biotechnology, Graduate School of Advanced Sciences of Matter, Hiroshima University, 1-3-1 Kagamiyama, Higashi-Hiroshima, Japan

Hiroko Kawasaki, Department of Molecular Biotechnology, Graduate School of Advanced Sciences of Matter, Hiroshima University, 1-3-1 Kagamiyama, Higashi-Hiroshima, Japan

L. Kenyon, AVRDC—The World Vegetable Center, Shanhua, Tainan 71499, Taiwan

Beom Soo Kim, Department of Chemical Engineering, Chungbuk National University, 12 Gaeshindong, Heungdeokgu, Cheongju, Chungbuk, Korea

Hak-Ryul Kim, Department of Animal Science and Biotechnology, Kyoungpook National University, Daegu, Republic of Korea

Takaaki Kiryu, Osaka Municipal Technical Research Institute, Osaka, Japan

Taro Kiso, Osaka Municipal Technical Research Institute, Osaka, Japan

Motomitsu Kitaoka, Enzyme Laboratory, National Food Research Institute, 2-1-12 Kannondai, Tsukuba, Ibaraki, Japan

Takashi Kuriki, Biochemical Research Laboratory, Ezaki Glico Co., Ltd., 4-6-5 Utajima, Nishiyodogawa-ku, Osaka, Japan

K.T. Lee, Department of Food Science and Technology, Chungnam National University, 220 Gung-Dong Yusung-Gu, Daejon, South Korea

R. Li, Plant Science, University of Kentucky, Lexington, KY, USA

Qing-Shan Li, Division of Applied Life Sciences, Graduate School of Agriculture, Kyoto University, Kitashirakwa-oiwakecho, Sakyo-ku, Kyoto 606-8502, Japan

Wen-Hsiung Li, Genomics Research Center, Academia Sinica, Taipei, Taiwan and Biodiversity Research Center, Academia Sinica, Taipei, Taiwan

Yebo Li, Department of Food, Agricultural, and Biological Engineering, The Ohio State University/Ohio Agricultural Research and Development Center, Wooster, OH, USA

Jiann-Tsyh Lin, Western Regional Research Center, ARS, USDA, Albany, CA, USA

Qin Liu, Agricultural Lipid Biotechnology Program, Department of Agricultural, Food and Nutritional Science, University of Alberta, 4-10 Agriculture/Forestry Centre, Edmonton, Alberta, Canada

Siqing Liu, Renewable Product Technology Research Unit, National Center for Agricultural Utilization Research, Agricultural Research Service, United States Department of Agriculture, 1815 N. University St., Peoria, IL61604, USA

Maria J. López, Departamento de Biología Aplicada, University of Almería, Almería, Spain

Thomas McKeon, Western Regional Research Center, United States Department of Agriculture, Albany, California, USA

Kazuo Miyashita, Faculty of Fisheries Sciences, Hokkaido University, 3-1-1 Mirato, Hakodate, Japan

Joaquín Moreno, Departamento de Biología Aplicada, University of Almería, Almería, Spain

Hiromi Murakami, Osaka Municipal Technical Research Institute, Osaka, Japan

Hiroshi Nabetani, Head of the Laboratory. Reaction and Separation Engineering Laboratory Food Engineering Division National Food Research Institute, National Agriculture and Food Research Organization, 2-1-12 Kan-nondai, Tsukuba, Ibaraki, Japan

Koji Nagao, Laboratory of Nutrition Biochemistry, Department of Applied Biochemistry and Food Science, Saga University, Saga, Japan

Mitsutoshi Nakajima, National Food Research Institute, NARO, Kan-nondai, Tsukuba, Ibaraki, Japan

Hirofumi Nakano, Osaka Municipal Technical Research Institute, 6-50, Morinomiya 1-chome, Joto-ku, Osaka, Japan

Maya Nanko, Department of Molecular Biotechnology, Graduate School of Advanced Sciences of Matter, Hiroshima University, 1-3-1 Kagamiyama, Higashi-Hiroshima, Japan

Nancy N. Nichols, Bioenergy Research Unit, NCAUR, USDA-ARS, Peoria, IL 61604, USA

Mamoru Nishimoto, National Food Research Institute, 2-1-12, Kannondai Tsukuba Ibaraki, Japan

Takahisa Nishimura, Biochemical Research Laboratory, Ezaki Glico Co., Ltd., 4-6-5 Utajima, Nishiyodogawa-ku, Osaka, Japan

Hiromi Nishiura, Biochemical Research Laboratory, Ezaki Glico Co., Ltd., 4-6-5 Utajima, Nishiyodogawa-ku, Osaka, Japan

Koji Nomura, Biochemical Research Laboratory, Ezaki Glico Co., Ltd., 4-6-5 Utajima, Nishiyodogawa-ku, Osaka, Japan

Jun Ogawa, Division of Applied Life Sciences, Graduate School of Agriculture, Kyoto University, Sakyo-ku, Kyoto, Japan

Kazuhisa Ono, Department of Molecular Biotechnology, Graduate School of Advanced Sciences of Matter, Hiroshima University, 1-3-1 Kagamiyama, Higashi-Hiroshima, Japan

Abrizah Othman, Malaysian Palm Oil Board, No. 6, Persiaran Institusi, Bandar Baru Bangi, Kajang, Selangor, Malaysia

Jocelyn A. Ozga, 4-10 Agriculture/Forestry Centre, Department of Agricultural, Food and Nutritional Science, University of Alberta, Edmonton, Alberta, Canada

Ramesh Patel, SLRPAssociates, Biotechnology Consulting Firm, Bridgewater, NJ, USA

Umi Salamah Ramli, Malaysian Palm Oil Board, No. 6, Persiaran Institusi, Bandar Baru Bangi, Kajang, Selangor, Malaysia

S. Rao, Plant Science, University of Kentucky, Lexington, KY, USA

Dennis M. Reinecke, Department of Agricultural, Food and Nutritional Science, University of Alberta, Edmonton, Alberta

J.S. Rhee, Department of Biological Sciences, Korea Advanced Institute of Science and Technology, 373-1, Guseong-dong, Yuseong-gu, Daejeon, Korea

Ravigadevi Sambanthamurthi, Malaysian Palm Oil Board, No. 6, Persiaran Institusi, Bandar Baru Bangi, Kajang, Selangor, Malaysia

Rolf D. Schmid, Institut für Technische Biochemie, Universität Stuttgart, Allmandring 31, Stuttgart, Germany

Ming-Che Shih, Agricultural Biotechnology Center, Academia Sinica, Taipei, Taiwan

Yuji Shimada, Osaka Municipal Technical Research Institute, Osaka, Japan. 1-6-50 Morinomiya, Joto-ku, Osaka, Japan

Sakayu Shimizu, Divison of Applied Life Sciences, Graduate School of Agriculture, Kyoto University, Kitashirakwa-oiwakecho, Sakyo-ku, Kyoto, Japan

Bungo Shirouchi, Laboratory of Nutrition Biochemistry, Department of Applied Biochemistry and Food Science, Saga University, Saga, Japan and Food Function and Labeling Program, National Institute of Health and Nutrition, 1-23-1 Toyama, Shinjuku-ku, Tokyo, Japan

Rodrigo M. P. Siloto, Agricultural Lipid Biotechnology Program, Department of Agricultural, Food and Nutritional Science, University of Alberta, 4-10 Agriculture/Forestry Centre, Edmonton, Alberta, Canada

J.K. Song, Chemical Biotechnology Research Center, Korea Research Institute of Chemical Technology, Yuseong-gu, Daejeon, Korea

Jae Yong Song, Department of Chemical Engineering, Chungbuk National University, Cheongju, Chungbuk, Republic of Korea

Lucia Slavikova, Department of Biochemical Technology, Faculty of Chemical and Food Technology, Slovak University of Technology, Radlinskeho 9, Bratislava, Slovak Republic

Kazuhisha Sugimoto, Biochemical Research Laboratory, Ezaki Glico Co., Ltd., Osaka, Japan

Masakazu Sugiyama, AminoScience Laboratories, Ajinomoto Co. Inc., 1-1 Suzukicho, Kawasakiku, Kawasakishi, Japan, 210-8681

Huang-Mo Sung, Department of Life Sciences, National Cheng Kung University, Tainan, Taiwan

Ahmed Tafesh, Institute of Applied Research Center (Affiliated with University of Haifa), The Galilee Society, Shefa-Amr, Israel

Koretaro Takahashi, Graduate School of Fisheries Science, Hokkaido University, 3-1-1 Minato, Hakodate, Japan

J.R. Thoguru, Plant Science, University of Kentucky, Lexington, KY, USA

Takayuki Tsukui, Faculty of Fisheries Sciences, Hokkaido University, 3-1-1 Minato, Hakodate, Japan

Vlada Urlacher, Institut für Technische Biochemie, Universität Stuttgart, Allmandring 31, Stuttgart, Germany

S. Venkatesan, AVRDC—The World Vegetable Center, P.O. Box 42, Shanhua, Tainan 71499, Taiwan

Mohd Basri Wahid, Malaysian Palm Oil Board, No. 6, Persiaran Institusi, Bandar Baru Bangi, Kajang, Selangor, Malaysia

Tzi-Yuan Wang, Genomics Research Center, Academia Sinica, Taipei, Taiwan

Yomi Watanabe, Osaka Municipal Technical Research Institute, 1-6-50 Morinomiya, Joto-ku, Osaka, Japan

Randall J. Weselake, Department of Agriculture, Food and Nutritional Sciences, University of Alberta, Edmonton, Alberta, Canada

M. Airanthi K. Widjaja-Adhi, Faculty of Fisheries Sciences, Hokkaido University, 3-1-1 Minato, Hakodate, Japan

Chi-Huey Wong, Department of Chemistry, The Scripps Research Institute, 10550 N. Torrey Pines Rd., La Jolla, CA 92037, USA

Shi Xiao, School of Biological Sciences, The University of HongKong, Hong Kong, China

Teruyoshi Yanagita, Department of Applied Biological Sciences, Saga University, Saga, Japan

Suk Hoo Yoon, Korea Food Research Institute, San 46-1, Baekhyun-Dong, Bundang-Ku, Songnam-Si, Kyunggi-Do, Korea

Abdul Masani Mat Yunus, Malaysian Palm Oil Board, No. 6, Persiaran Institusi, Bandar Baru Bangi, Kajang, Selangor, Malaysia

Jitao Zou, Plant Biotechnology Institute, National Research Council Canada, 110 Gymnasium Place, Saskatoon, Canada

Section I

Improvement of Agronomic and Microbial Traits

1

Insights Into the Structure and Function of Acyl-CoA: Diacylglycerol Acyltransferase

Rodrigo M.P. Siloto, Qin Liu, and Randall J. Weselake

Agricultural Lipid Biotechnology Program, Department of Agricultural, Food and Nutritional Science, University of Alberta, 4-10 Agriculture/Forestry Centre, Edmonton, Alberta, Canada T6G 2P5

Xiaohua He and Thomas McKeon

Western Regional Research Center, United States Department of Agriculture, Albany, California 94710, USA

1.1 Introduction

Production of vegetable oils has been recognized as a rapidly developing field in plant biotechnology that goes beyond food-based applications. Many kinds of vegetable oils are used in soaps and cosmetics or converted to oleochemicals that are extensively used to replace petrochemicals in paints, plastics, fuels, and lubricants. The demand for biodegradable chemicals applied to industrial products has been increasing, and therefore a boost in the production of vegetable oils and fats is needed. Biotechnological approaches including traditional plant breeding and direct genome modification through genetic engineering are crucial tools to increase seed oil production without extending the area of crop cultivation, which has a direct impact on deforestation and competition with food production. Moreover, even a diminutive increase in seed oil content reflects in considerable profitability. Despite the unprecedented advances derived from molecular genetics and genomics research on the biochemical pathways of plant lipid metabolism in the last decade, the mechanisms regulating seed oil content are not fully understood. Many aspects of key enzymes are not yet determined even in model plants such as Arabidopsis thaliana (Hildebrand et al., 2008). For example, recent studies focusing on intracellular trafficking indicated that compartmentalization of enzyme activities within the endoplasmic reticulum (ER) membrane represents an additional mechanism adopted by plant cells to control oil production and may be essential for channeling of particular fatty acids into storage lipids (Dyer and Mullen, 2008).

Nevertheless, manipulation of genes involved in storage lipid biosynthesis has been used to increase accumulation of seed triacylglycerol (TAG), the main component of vegetable oils (Weselake, 2002). It was recently demonstrated that overexpression of plant and fungi genes encoding acyl-CoA:diacylglycerol acyltransferase (DGAT, EC 2.3.1.20), which catalyzes the final assembly of TAG, resulted in small but significant increases in seed oil content in canola and soybean tested under field conditions (Lardizabal et al., 2008; Weselake et al., 2008). Indeed, the level of DGAT activity in developing seeds seems to have a direct effect on the accumulation of TAG (Perry and Harwood, 1993; Cahoon et al., 2007). Surprisingly, little is known about the molecular mechanisms governing DGAT activity. The most basic information about structure and function of this enzyme is essential for rational designs to increase its performance in oilseeds and have a direct reflection in seed oil content. In view of the biotechnological importance of DGATs from plants and fungi, we summarize some of the structural and functional aspects of these enzymes with particular attention to membrane topology, functional polypeptide motifs, and subcellular localization. We use in silico approaches to compare the findings obtained with related enzymes in animals and prokaryotes.

1.2 Discovery of DGAT

The first proceedings reporting DGAT activity date from the 1950s (Weiss and Kennedy, 1956; Weiss et al., 1960), but the genes encoding DGATs were not isolated until the late 1990s. The first DGAT cDNA was cloned by taking advantage of homology between an expressed sequence tag (EST) and an acyl-CoA:cholesterol acyltransferase (ACAT, EC 2.3.1.26), a related enzyme previously isolated by a complementation assay of mammalian cells devoid of cholesterol ester biosynthesis (Chang et al., 1993). The mouse (Mus musculus) DGAT gene isolated in 1998 encodes a protein, here referred to as MmDGAT1 that is 20% identical to mouse ACAT with the most conserved regions on the C-terminus portion of the enzyme (Cases et al., 1998). A plant DGAT gene was consequently isolated through the characterization of the locus TAG1 in an A. thaliana EMS-induced mutant (AS11) with altered seed fatty acid composition and decreased DGAT activity (Katavic et al., 1995). The locus TAG1 contains a 3.4-kb gene encoding a polypeptide showing 41% identity with MmDGAT1 (Zou et al., 1999). The polypeptide encoded by TAG1 (AtDGAT1) exhibits DGAT activity when expressed in yeast and can complement DGAT function in AS11 (Jako et al., 2001). DGAT genes from fungi were identified through protein purification, an approach that was previously not successful with other DGATs, perhaps because of their membrane association. Polypeptides exhibiting DGAT activity were purified from lipid bodies of Umbelopsis ramanniana, formerly known as Mortierella ramanniana (Lardizabal et al., 2001). These DGATs shared little or apparently no homology with the previous DGAT genes, and therefore were classified as DGAT2. Curiously, genes homologous to DGAT1 have not been found in fungi genomes, although it has been suggested that yeast ACATs (ARE1 and ARE2 in Saccharomyces cerevisiae) represent DGAT1 orthologs in these organisms because they also display minor DGAT activity (Yen et al., 2008).

Several lines of evidence suggest that DGAT1 belongs to a class of enzymes with acyl-CoA transferase activity, which can utilize different acceptors in addition to diacylglycerols. For example, MmDGAT1 also possesses acyl-CoA:retinol acyltransferase (ARAT, EC 2.6.1.57) activity (Yen et al., 2005), while an A. thaliana acyl-CoA:fatty alcohol acyltransferase (wax ester synthase, WSD1) also displays DGAT activity in vitro (Li et al., 2008). In the case of DGAT2, a similar scenario is observed. In animals, DGAT2 belongs to a gene family with seven members in humans (Cases et al., 2001). Three of these genes encode polypeptides with acyl-CoA monoacylglycerol acyltransferase (MGAT, EC 2.3.1.22) activity (Yen et al., 2002; Yen and Farese, 2003; Cheng et al., 2003). Two additional members display acyl-CoA:wax alcohol acyltransferase (AWAT, EC 2.3.1.75) activity, which is analogous to WSD1 (Turkish et al., 2005).

Orthologs of DGAT1 and DGAT2 have been identified through DNA homology in many other organisms and are widely distributed in eukaryotes. Currently, a relatively wide collection of DGAT genes is available which facilitates more detailed studies of enzyme structure and function through bioinformatic approaches. Many of these genes have been functionally characterized in recombinant systems as described in Table 1.1.

Table 1.1 Eukaryotic DGATs Functionally Tested in Recombinant Organisms

In prokaryotes, a bifunctional WS/DGAT was identified in Acinetobacter calcoaceticus (Kalscheuer and Steinbuchel, 2003). WS/DGAT has no sequence similarity to DGAT1, DGAT2, or any of the related acyltransferases from eukaryotes. Another nonhomologous DGAT, referred to as AhDGAT, was characterized in peanuts (Saha et al., 2006). Unlike other eukaryote enzymes, AhDGAT was purified from the soluble fraction of developing peanuts. Biosynthesis of TAG in the cytosol has been previously reported in a 10S multienzyme complex from the oleaginous yeast Rhodotorula glutinis (Gangar et al., 2001). Whether this soluble yeast DGAT and AhDGAT compose a novel class of DGATs is yet to be demonstrated.

1.3 Membrane Topological Organization of DGATs

The pattern in which a protein transverses the membrane bilayer is essential for elucidating the dynamics of the protein structure. DGAT1 and DGAT2 contain hydrophobic segments that are generally believed to constitute transmembrane domains (Fig. 1.1). DGAT1 displays more hydrophobic segments than DGAT2, which indicates a different topology and may relate to different physiological roles in TAG biosynthesis (Yen et al., 2008). Few experimental studies on DGAT topological organization in plants and yeast are available, and therefore we will mainly rely on in silico approaches to predict transmembrane segments and the orientation in the membrane bilayer.

Figure 1.1 Kyte–Doolittle hydropathy plots of DGATs. Plots were generated by the method of Kyte and Doolittle (1982) using a window size of 19. Cutoff value (line) is 1.8 and peaks with score greater than 1.8 indicate possible transmembrane regions.

A variety of web-based tools are available for predicting the topology of membrane proteins. Since only a few membrane proteins from bacteria are known to be beta-barrel shaped so far, the prediction algorithms are mostly developed for alpha-helical membrane proteins. Generally, five types of techniques have been used in these programs: hydrophobicity analysis combined with the positive inside rule (eg. TMpred and SOSUI), multiple sequence alignment (eg. ConPredII and TOPCONS), model-recognition approach (eg. MEMSTAT3, TMHMM, and HMMTOP), and support vector machine technique (eg. SVMtm) (Persson, 2006). An evaluation of the reliability of these methods indicated that a consensus prediction and model-based methods are best performing (Moller et al., 2001; Ikeda et al., 2002). The application of these algorithms for the prediction of TM domains in DGAT1 is described in Table 1.2 using AtDGAT1 and MmDGAT1 as models. For AtDGAT1, nine of the ten putative transmembrane domains are highly conserved among most of the prediction results except for the domains at 276–299 and 314–337 of AtDGAT1 and 251–276 and 285–312 of MmDGAT1 (highlighted TM5 and TM6 in Table 1.2). A model of nine-membrane-spanning topology agrees with our initial study on DGAT1 from Brassica napus (Foroud, 2005). In this work, protease mapping data showed that the region between 276 and 299 in BnDGAT1 (corresponding to TM5) is in the cytosol, in agreement with most of the prediction algorithms described in Table 1.2. Recent studies on DGAT1 from Vernicia fordii (tung tree) and B. napus indicated that the N-terminus faces the cytosolic side (Shockey et al., 2006; Weselake et al., 2006) as predicted by most algorithms in Table 1.2. The interaction of the N-terminus with lipid substrates in the cytoplasm may lead to a regulatory role of N-terminal region (Siloto et al., 2008) and there are several lines of evidence not only from B. napus DGAT1 but from mammalian DGAT1 and ACAT1 that favor this hypothesis (Cheng et al., 2001; Yu et al., 1999; Weselake et al., 2006). According to the work on VfDGAT1, the C-terminus of DGAT1 is also proved to orient toward cytosolic side, indicating an even number of membrane-spanning regions. This result disagrees with a nine-transmembrane topology model, and therefore further experimental testing will be required to examine the hypothesis of eight transmembrane domains.

Table 1.2 Prediction Results for Transmembrane Domains in DGAT1

Compared to DGAT1, DGAT2 is less hydrophobic, having a lower number of transmembrane domains and therefore a less intricate topology. The membrane topology of MmDGAT2 was experimentally determined revealing two transmembrane domains that are closely associated or a single hydrophobic domain embedded in the membrane bilayer (Stone et al., 2006). The first transmembrane domain (TM1) of MmDGAT2 and ScDGAT2 was ubiquitously predicted, but the second (TM2) was identified by only a few algorithms (Table 1.3). Since the homology of DGAT2 from different organisms is lower than that of DGAT1, it is possible that ScDGAT2, which has a distinct hydropathy plot, could have a different topology compared with other fungi DGAT2s. This could be demonstrated by the prediction results of Schizosaccharomyces pombe SpDGAT2 (Table 1.3). Interestingly, the prediction of N-terminus orientation seems to be related to the length of the predicted N-terminal tail. DGAT2s with putative long tails are intended to face toward the cytosol, which agrees with work on VfDGAT2 and MmDGAT2 (Shockey et al., 2006; Stone et al., 2006). The same conclusion, however, cannot be made for DGAT2s with short tails.

Table 1.3 Prediction Results for Transmembrane Domains in DGAT2

1.4 Alignment of DGAT1 Polypeptides

DGAT1 polypeptides are typically characterized by a hydrophilic N-terminus sequence followed by a number of hydrophobic stretches constituting potential transmembrane domains as previously discussed. The total number of predicted transmembrane domains in DGAT1 can vary according to the sequence and the algorithm used as shown above. When the sequences are aligned, however, many of these potential transmembrane domains are found in the same positions in most DGAT1 (Fig. 1.2). The first four transmembrane domains on the first half of the sequences and the last three transmembrane domains on the C-terminus are separated by short polar loops. Between these groups are two possible membrane-spanning regions that are separated by longer hydrophilic stretches. Here we will consider these nine potential transmembrane domains as landmarks to describe conserved motifs in DGAT1, acknowledging, however, that an experimental approach is required to verify these assumptions. We will also use the sequence of A. thaliana DGAT1 to describe the exact position of each motif.

Figure 1.2 Alignment of transmembrane domains in DGAT1. The putative transmembrane domains of DGAT1 polypeptides from 12 animal and 18 plant organisms were predicted and the polypeptides were aligned. The identity of the alignment is graphed on the top using a window size of 6. The arrows denote the predicted transmembrane domains. The thick lines represent the sequence of each DGAT1, and the thin lines represent the gaps generated by the alignment. The picture was generated with Geneious Pro 4.6.0 and optimized manually. The transmembrane domains were predicted with transmembrane hidden Markov model (TMHMM). Accession numbers for the DGAT1 polypeptides are: AtDGAT1, NM_127503; AaDGAT1, XP_001658299; BnDGAT1, AAD45536; CeDGAT1, CAB07399; DmDGAT1, AAL78365; DrDGAT1, NP_956024; EaDGAT1, AAV31083; GmDGAT1, AAS78662; HsDGAT1, NP_036211; JcDGAT1, ABB84383; MdDGAT1, XP_001371565; MmDGAT1, NP_034176; MtDGAT1, ABN09107; NtDGAT1, AAF19345; NvDGAT1, XP_001639351; OeDGAT1, AAS01606; OsDGAT1, BAD53762; PfDGAT1, AAG23696; PpDGAT1, XP_001770929; PtDGAT1, XP_002330510; RcDGAT1, AAR11479; RnDGAT1, BAC43739; SsDGAT1, NP_999216; TaDGAT1, XP_002112025; TgDGAT1, AAP94209; TmDGAT1, AAM03340; VfDGAT1, ABC94471; VgDGAT1, ABV21945; VvDGAT1, CAN80418; ZmDGAT1, ABV91586.

An overview of the DGAT1 alignment from 30 different organisms indicates several conserved regions with about 7% of identical residues among plant and animal sequences. The hydrophilic N-terminus is composed of an average of 115 and 80 residues in plants and animals, respectively and is the least conserved region in DGAT1. An alignment of the N-terminal portion of DGAT1 from a broad range of organisms revealed a cluster of arginines in the first 30 residues (Fig. 1.3). The region comprising 20 positions preceding the first hydrophobic domain is also conserved and contains the motifs PAHRXXXESPLSSDAIFXQ and SLFSXXSGFXN, which are conserved in plants and animals, respectively. Other divergences discriminating DGAT1 from plants and animals include a serine at position 131 of AtDGAT1 conserved in plants and absent in animal DGAT1, and the motif WVXRQ in plants, corresponding to FL(L/I)(R/K)R in animals. These differences can be also observed in more ancient organisms such as Toxoplasma gondii and Physcomitrella patens. The long loop between the fourth and fifth transmembrane domains (between positions 260 and 278 of AtDGAT1) shows remarkable variability among all DGAT1s. Following this region lies the most conserved uninterrupted sequence of DGAT1 comprising the motifs PTLCYQXSYPR in plants and PTLCYEXXFPR in animals, preceding the fifth predicted transmembrane domain between positions 292 and 297 of AtDGAT1.

Figure 1.3 Alignment of the N-terminus polypeptide sequence of DGAT1 from plants and animals. Gray shades denote the polarity of blocks of conserved residues. The position corresponding to the end of the first exon in plants is indicated.

1.5 Alignment of DGAT2 Polypeptides

DGAT2 polypeptides, in comparison to DGAT1, display fewer potential transmembrane domains and higher sequence divergence. An alignment of DGAT2 sequences from 20 organisms, covering plants and animals previously described in DGAT1, indicate approximately 5% of identical residues. Inclusion of 16 fungi sequences in this group decreases the identity to only 2.3%. A higher divergence might pose difficulties for identification of novel members of DGAT2 through sequence homology.

At least one transmembrane domain can be predicted for every DGAT2, but usually two transmembrane domains are conserved in the N-terminus portion and separated by a small loop (Fig. 1.4). This hydrophobic region is definitely very important because its removal results in lack of activity in ScDGAT2 (unpublished). An experimental approach indicated that the only membrane-spanning region in MmDGAT2 is composed of two transmembrane domains separated by a small loop that could be also interpreted as a single hydrophobic region embedded in the membrane (Stone et al., 2006). This region will be used as a landmark and the positions of conserved motifs also will be indicated in the UrDGAT2a polypeptide from U. ramanniana. The N-terminus portion preceding the transmembrane domains is quite variable in length and is usually smaller in animals and plants (with 38 and 30 residues in average, respectively) when compared with fungi (with 100 residues in average). The most conserved region in DGAT2 encompasses the motif RXGFX(K/R)XAXXXGXX(L/V)VPXXXFG(E/Q) located approximately 150 residues after the second transmembrane domain (positions 259–281 of UrDGAT2a). Other conserved residues are the motif GGXXE (positions 204–208 in UrDGAT2a) and a phenylalanine, an arginine, and a proline in positions 164, 170, and 293 of UrDGAT2a, respectively. In addition, the motif YXXXXXHPHG is conserved in sequences from animals and fungi (positions 121–129 of UrDGAT2a) corresponding to YXXXXXEPHS/G in plants. Preceding this motif is situated one of the most striking divergences in DGAT2 alignment, a hydrophilic segment of approximately 41 residues present in sequences from some fungi but absent in plants and animals. This region, corresponding to positions 144–185 of ScDGAT2, is also found as a much larger segment (158 residues) in Yarrowia lipolytica DGAT2. This hydrophilic segment, although nonessential, was demonstrated to modulate the enzyme activity of ScDGAT2 (unpublished results). Because this segment precedes a highly conserved motif, it is possible that it might represent a specialized function in DGAT2 from certain fungi.

Figure 1.4 Alignment of transmembrane domains in DGAT2. The putative transmembrane domains of DGAT2 polypeptides from 18 fungi, 5 animals, and 11 plants were predicted and the polypeptides were aligned. The identity of the alignment is graphed on the top using a window size of 6. The arrows denote the predicted transmembrane domains. The thick lines represent the sequence of each DGAT2 polypeptide, and the thin lines represent the gaps generated by the alignment. The picture was generated as described for DGAT1. Accession numbers for the DGAT2 polypeptides are: AcDGAT2, XP_001540241; AdDGAT2, XP_001273210; AnDGAT2, CAK46407; AoDGAT2, XP_001822244; AtDGAT2, NP_566952; BfDGAT2, XP_002208225; BtDGAT2, CAD58968; CeDGAT2, CAB04533; CeDGAT2b, AAB04969; CiDGAT2, XP_001240299; CnDGAT2, EAL20089; CrDGAT2, XP_001693189; DdDGAT2, XP_635762; GzDGAT2, XP_381525; HsDGAT2, AAK84176; LbDGAT2, EDR14458; MgDGAT2, XP_368741; MmDGAT2, AAK84175; MtDGAT2, ACJ84867; NcDGAT2, CAE76475; NfDGAT2, XP_001261291; OsDGAT2, NP_001057530; OtDGAT2, CAL58088; PmDGAT2, XP_002146410; PnDGAT2, EAT89076; PpDGAT2, XP_001777726; PtDGAT2, XP_002317635; RcDGAT2, AAY16324; ScDGAT2, NP_014888; SpDGAT2, XP_001713160; TsDGAT2, EED21737; UmDGAT2, XP_760084; UrDGAT2a, AAK84179; UrDGAT2b, AAK84180; VfDGAT2, ABC94474; VvDGAT2, CAO68497; ZmDGAT2, ACG38122.

1.6 Structure of DGAT Genes

The architecture of genes encoding DGAT is largely available from whole-genome sequence databases or, as in the case of V. fordii, from sequencing of the respective genomic regions. In mammals, genes encoding DGAT1 share a similar architecture of 17 exons mostly grouped in the 3′ portion. A DGAT1 representative from invertebrates (Caenorhabditis elegans), however, shows an unrelated distribution with only seven exons (Fig. 1.5A). In plants, DGAT1 genes from A. thaliana, M. truncatula, Z. mays, and V. fordii are composed of 16 exons, while DGAT1 from O. sativa contains 14 exons (Fig. 1.5B). The first exon of plant DGAT1 genes comprises the largest coding sequence and encodes the hydrophilic N-terminus. Curiously, the last codon from the first exon of these genes encodes a glutamine in the same position of the alignment (motif IFXQ), denoting the end of the hydrophilic N-terminus and start of the first predicted membrane-spanning region (Fig. 1.3). The hydrophilic N-terminus is the most variable sequence of DGAT1 polypeptides, and therefore it is possible that segregation of this sequence in the first exon might have been used as an evolutionary mechanism to delimit variability in this region of the gene. This pattern was not observed in DGAT1 sequences from animals. DGAT2 genes show a structure that is dissimilar to that of DGAT1. Mammalian DGAT2 genes share a common architecture with eight exons while the gene from C. elegans has only two exons (Fig. 1.5C). In plants DGAT2 genes have eight exons in A. thaliana and ten exons in V. fordii and O. sativa (Fig. 1.5D).

Figure 1.5 Architecture of DGAT genes. (A) DGAT1 from animals, (B) DGAT1 from plants, (C) DGAT2 from animals, and (D) DGAT2 from plants. The genomic sequences of each DGAT are represented by black bars. The arrows correspond to the regions comprising the coding region of the mRNA. The numbers correspond to the nucleotide positions. Accession numbers are: HsDGAT1, AC_000140.1; MmDGAT1, NC_000081.5; SsDGAT1, AY116586.1; CeDGAT1, NC_003283.9; VfDGAT1, DQ356679.1; MtDGAT1, AC174465.2; ZmDGAT1, AM433916.2; OsDGAT1, AP008212.1; AtDGAT1, NC_003071.4; HsDGAT2, NC_000011.8; BtDGAT2, NC_007313.3; MmDGAT2, NC_000073.5; CeDGAT2, Z81557.1; VfDGAT2, DQ356681.1; OsDGAT2, AP004757.3 and AtDGAT2, NC_003074.5.

1.7 Functional Motifs in DGAT1

Most of the information available on the structure and function of DGATs is derived from comparisons of homologous enzymes. Alignments of polypeptide sequences encoding acyl-CoA-dependent acyltransferases from diverse organisms indicated a conserved histidine and an aspartic acid in the configuration HXXXXD. Substitution of the conserved histidine in the bifunctional enzyme 2-acyl-glycerophosphoethanolamine acyltransferase/acyl–acyl carrier protein synthase (Aas, EC 2.3.1.40 and 6.2.1.20, respectively) resulted in lack of acyltransferase activity (Heath and Rock, 1998). Substitution of the aspartic acid residue also resulted in significantly less activity. It was suggested that the histidine operates as a general base to abstract the proton from the hydroxyl group of the sn-1 glycerol-3-phosphate, facilitating nucleophilic attack on the thioester bond of acyl-CoA. The aspartic acid would work in a charge relay system to increase the nucleophilicity of the hydroxyl group. This mechanism could be used by other acyltransferases, including DGAT. In fact a similar motif (HHXXXDG) is conserved in DGATs from prokaryotes (Daniel et al., 2004). In eukaryotic DGAT1, the motif HXXXD can be found closely after the fourth predicted transmembrane domain in DGAT1 from plants (positions 257–261 of AtDGAT1). Similarly, the motif HXXXXD is found in a region preceding the fifth predicted transmembrane domain of a few plants such as A. thaliana, B. napus, R. communis, and V. fordii (positions 342–347 of AtDGAT1) (Fig. 1.6). These motifs, however, are not conserved in animals and therefore might not compose the catalytic site of DGATs. Jako et al. (2001) identified the consensus sequence N(S/A/G)R(L/V)(I/F/A)(I/L)EN(L/V) in AtDGAT1 and proposed that the invariant arginine and glutamic acid on positions 149 and 153 could have functions analogous to those of histidine and aspartic acid residues, respectively. This region is highly conserved in all organisms including more ancient eukaryotes (T. gondii and P. patens) (Fig. 1.6). These residues are present in the interface between a putative transmembrane domain and the adjacent hydrophilic loop, which would create an amphipathic environment for the substrates of DGAT. Moreover, DGAT1 is recognized as a member of a large protein family of membrane-bound O-acyltransferases known as MBOAT (NCBI domain ID pfam03062; Hofmann, 2000). Other members of the MBOAT family catalyze O-acylation reactions transferring acyl chains onto hydroxyl or thiol groups of lipids and proteins. For example, ACAT transfers an acyl chain from acyl-CoA to cholesterol, forming cholesteryl esters (Chang et al., 1993) while skinny hedgehog (ski) protein transfers a palmitoyl group onto cysteine residues of other proteins (Chamoun et al., 2001). The MBOAT family is characterized by a hydrophobic region (positions 234–509 of AtDGAT1) that contains a conserved asparagine (position 410 in AtDGAT1) and histidine (position 447 in AtDGAT1) (Fig. 1.6). It has been proposed that these residues could be involved in the catalytic activity. For example, this conserved histidine has been demonstrated to be a key residue for human ACAT1 activity (Guo et al., 2005). Whether any of these regions contribute to the catalytic site of DGAT1 is yet to be experimentally tested. Interestingly, sn-1 glycerol 3-phosphate acyltransferase (GPAT, EC 2.3.1.15) and lysophosphatic acid acyl transferase (LPAAT, EC 2.3.1.51), which are also membrane-bound O-acyltransferases catalyzing the first two acylation steps of TAG biosynthesis, are not classified as MBOAT members, suggesting that these enzymes might not share similar catalytic sites. It is also possible that these residues could act as supplementary catalytic sites being involved in other enzyme activities besides DGAT, such as ARAT and ACAT.

Figure 1.6 Alignment of putative active sites in DGAT1. A scheme MmDGAT1and AtDGAT1 is described on the top with the position of the MBOAT motif. The arrows in this scheme represent the predicted transmembrane domains. The thick lines represent the sequence of each DGAT1 polypeptide, and the thin lines represent the gaps generated though the alignment as previously shown. The vertical boxes contain the amino acid sequences for different DGATs indicated on the left. The arrows on these boxes indicate the position of conserved residues discussed in the text. Accession numbers for the DGAT polypeptides are the same as in Figure 1.2.

Other putative active sites in DGAT1 include the substrate binding sites. Sequences of DGAT1 from several plants indicate the presence of a putative diacylglycerol/phorbol ester binding motif that is apparently absent in ACATs (Zou et al., 1999; Nykiforuk et al., 2002; Xu et al., 2008). Phorbol esters such as phorbol-12-myristate-13-acetate (PMA) are commonly known to mimic diacylglycerols. The putative diacylglycerol/phorbol ester binding motif present in the positions 414 and 424 of AtDGAT1 forms the consensus HXXXXRHXXXP in DGAT1 from plants and animals. Xu et al. (2008) demonstrated that substitution of a phenylalanine by an arginine in position 439 of TmDGAT1 that is 16 positions after the predicted motif resulted in loss of DGAT activity. This could be a result of alterations in DAG interaction with DGAT. But, because this phenylalanine is positioned at a predicted transmembrane domain, substitution by a charged residue could also have structural implications. Acyl-CoA has been shown to interact with a recombinant N-terminal segment of BnDGAT1 and MmDGAT1 (Weselake et al., 2000, 2006; Siloto et al., 2008). The N-terminus sequence is highly variable, except for a region of 20 residues preceding the first hydrophobic domain, which shows remarkable conservation among plants and animals. Many of these variations, however, represent amino acid residues with similar properties, which could explain the acyl-CoA binding properties of DGAT1 from B. napus and M. musculus. Several lines of evidence suggest that acyl-CoA interaction with the hydrophilic N-terminus of DGAT1 regulates this enzyme allosterically. First, there is positive cooperativity exhibited for binding of 22:1-CoA in mouse and canola DGAT1 (Weselake et al., 2000; Siloto et al., 2008). Second, enzymes that are allosterically regulated often form multimeric complexes to achieve cooperativity and the N-terminus of DGAT1 assists in the formation of dimers and tetramers as demonstrated for BnDGAT1 and HsDGAT1, respectively (Weselake et al., 2006; Cheng et al., 2001). For example, ACAT1 self-associates through the N-terminus, which also plays a regulatory role in this enzyme (Guo et al., 2001; Yu et al., 2002). Third, the acyl-CoA binding motif is not essential for enzyme activity because the removal of the N-terminus of RcDGAT1 results in a polypeptide with substantial enzyme activity, indicating that this is not the exclusive region to interact with acyl-CoA (unpublished data). Indeed, the fourth conserved block in GPATs and LPAATs, as described by Lewin et al. (1999), contains an invariant proline that has been proposed to participate in acyl-CoA binding. This proline was identified in plant DGAT1 polypeptides on the third predicted transmembrane domain corresponding to position 224 of AtDGAT1 and is in fact conserved in DGAT1 from all organisms. Substitution of this proline with an arginine in TmDGAT1 abolished DGAT activity, corroborating with the idea that this residue has a functional role (Xu et al., 2008). Another possible acyl-CoA binding site was proposed to be closely associated with the motif FYXDWWN in ACATs (Yen et al., 2008). This motif is present in DGAT1 and shows remarkable conservation with exception to CeDGAT1, where the second tryptophan is substituted by a phenylalanine. This motif is located on the loop preceding the third last putative transmembrane domain, relatively distant from the proline residue previously discussed, but near the asparagine residue conserved in MBOAT members. The paired tryptophans in this motif are a rare combination and have been previously demonstrated to participate in cholesterol binding. Guo et al. (2001) demonstrated, however, that substitution of the conserved tyrosine by alanine in yeast ACAT1 resulted in decreased affinity for acyl-CoA. Substitution of this same residue in TmDGAT1 (Y392A) resulted in decreased enzyme activity while a double mutation in tyrosine and tryptophan (Y392G/W395G) completely abolished enzyme activity (Xu et al., 2008).

Other putative functional domains predicted in DGAT1 include a leucine zipper and phosphorylation sites, although it is not yet clear whether these regions are important in the function, structure, or regulation of DGAT1. A putative leucine zipper motif was described in several DGAT1 from plants (Bouvier-Nave et al., 2000a; Nykiforuk et al., 2002). For example, in AtDGAT1 polypeptides five leucines (L222, L229, L236, L243, and L250) are consecutively spaced by six residues forming a classic leucine zipper (Hobbs et al., 1999). This leucine zipper, which might mediate interactions with other proteins, is present in a number of DGAT1 from plants but not from animals. Several studies indicated the presence of multiple potential phosphorylation sites in DGAT1 (Hobbs et al., 1999; Nykiforuk et al., 2002; He et al., 2004). Some of these sites are conserved in plant DGATs, such as the protein kinase C sites in the loop between the first and second transmembrane domains (positions 169–171 and 172–175 of AtDGAT1) and the casein kinase II sites (positions 254–257 and 403–406 of AtDGAT1). In addition, a tyrosine kinase site (positions 386–393 of AtDGAT1) is conserved in DGAT1 from plants and animals. This site overlaps with the FYXDWWN motif previously discussed as a putative acyl-CoA binding site. Although substitution of the conserved tyrosine by alanine in yeast ACAT homologue resulted in lower affinity to acyl-CoA, phosphorylation could not be directly detected (Guo et al., 2001). Regulation of DGAT1 activity through phosphorylation is compelling not only because this is a common mechanism to control enzyme activity in eukaryotes but also because DGAT can scavenge DAG, an important molecule involved in phosphorylation signaling cascades (Carrasco and Merida, 2007). For example, the affinity of DAG to C1 domains of DAG kinase is modified by phosphorylation of residues situated close to this motif (Thuille et al., 2005). In addition, the fact that DGAT is expressed in vegetative tissues suggests that it can have additional roles beyond oil biosynthesis in seeds (Lu et al., 2003). Substitution of serine at position 168 in RcDGAT1 that corresponds to a protein kinase C site previously described resulted in a significant decrease in the enzyme activity (unpublished).

1.8 Functional Motifs in DGAT2

The motifs previously described for DGAT1 cannot be found in DGAT2 sequences likely due to the little homology between DGAT1 and DGAT2. In fact, little is known about functional motifs of DGAT2. Stone et al. (2006) identified the conserved motif HPHG in positions 161–164 of MmDGAT2 as an important region for DGAT activity. Substitution of these residues, forming the sequences APHG, HGHG, HPAG, and AGAG, resulted in a significant reduction of enzyme activity. More specifically, the histidine at position 163 of MmDGAT2 appeared to play a more important role for the enzyme function, which agrees with our mutagenesis work on ScDGAT2 (unpublished). This region is conserved in animal and fungi DGAT2, but in plants this motif is found as EPHS/G. Substitution of the glutamic acid by a histidine residue in plant DGAT2 did not result in an appreciable effect, but replacement of the motif HPHG in ScDGAT2 with residues EPHS found in plant DGAT2 resulted in loss of enzyme activity (unpublished). This indicates an important divergence on the structure/function of DGAT2 from fungi and plants. In addition, the motif FLXLXXXn (n indicates a nonpolar residue) was indicated as a putative neutral lipid binding domain in MmDGAT2 (Stone et al., 2006). Substitution of phenylalanine (position 80) and leucine (position 81) residues by alanine residues resulted in decreased DGAT activity. Substitution of the leucine in position 83 by an alanine resulted in lack of activity. This motif, present in the first predicted transmembrane domain of MmDGAT2 (positions 80–87), is conserved in vertebrate DGAT2 but not in plants or fungi orthologs. Substitution of the corresponding phenylalanine and leucine (positions 71 and 73) in ScDGAT2 results in a decrease of approximately 50% of the wild-type activity (unpublished). This same motif contains the putative membrane lipid attachment LGVAC found in prokaryotes through the interaction between the sulfhydryl group of a cysteine residue (position 87 of MmDGAT2) and DAG. Substitution of this cysteine by a serine in MmDGAT2 did not reduce DGAT activity, indicating that it does not function as a lipid attachment site. In fact, substitution of all cysteine residues in ScDGAT2 by alanine residues did not disrupt DGAT activity, indicating that this mechanism of DAG interaction is not present or at least essential in this enzyme (unpublished data). In addition, substitutions on the conserved motif YFP located close to the transmembrane domains (positions 104–106 of UrDGAT2A) resulted in significant decreases in the enzyme activity.

1.9 Subcellular Localization of DGATs

To better elucidate the role of DGATs in cellular processes, their spatial location has been studied in different plants. In numerous earlier studies, DGAT location has been a subject of discrepancy whether it is associated with ER or oil bodies (Lung and Weselake, 2006). This debate could be the result of technique limitations because the general approach used in these studies was subcellular fractionation combined with enzyme assay in which cross-contamination can occur. For instance, in the study of germinating soybean cotyledon, the purified oil bodies also exhibited activities for ER markers (Settlage et al., 1995). This could be explained by association between oil bodies and ER (Cao and Huang, 1986; Settlage et al., 1995). Lacey and Hills (1996) applied different organelle markers to rule out the possible contamination in the assay and clearly demonstrated that B. napus DGAT is associated with ER. Similarly, Cao and Huang (1986) were able to localize maize DGAT in the rough ER (RER) by taking advantages of protein markers as well as the attachment of RER with polysomes in the presence of Mg2+ during fractionation. Actually the ER is regarded as the main site for TAG synthesis, and microsomal fractions from developing seeds of many plants as well as plant cultured cells have been extensively utilized for enzyme assays (Browse and Somerville, 1991; Weselake, 2002). Using more dependable techniques such as green fluorescent protein (GFP)-tagging and immunofluorescence, Shockey et al. (2006) have demonstrated that tung tree DGAT1 and DGAT2 are localized in the ER. Localization of both DGATs is dependent on a C-terminal ER retrieval motif. In VfDGAT1, the ER retrieval sequence YYHDL is part of the motif LLYYHDXMN conserved in all plant DGAT1. The ER retrieval domain in VfDGAT2 comprises the sequence LKLEI, where the two leucines are conserved in other DGAT2 sequences. Removal of the corresponding regions through C-terminus truncations in RcDGAT1 and ScDGAT2 resulted in decreased activity and decreased protein stability, respectively, indicating the importance of the C-terminus portion for both DGATs (unpublished). Interestingly, VfDGAT1 and VfDGAT2 do not colocalize in the ER, and therefore it is plausible that these polypeptides have distinct interactions with other proteins in the ER membrane. Mounting evidence based on studies with animals and plants indicate that DGAT1 and DGAT2, although catalyzing the same enzyme activity, have distinct physiological functions (Yen et al., 2008; Shokey et al., 2006; Burgal et al., 2008). In addition to the ER, DGAT activity was also found in chloroplasts of spinach leaves (Martin and Wilson, 1984) and more recently, Kaup et al. (2002), identified DGAT1 in the chloroplasts of senescing Arabidopsis leaves through immunoblotting. The mechanisms by which AtDGAT1 is transported to the chloroplast are yet to be determined.

In yeast, biochemical studies with S. cerevisiae indicated that DGAT activity is mainly in lipid droplets (Sorger and Daum, 2002). Indeed, DGAT2 in U. rammaniana was purified from the lipid particle fractions (Lardizabal et al., 2001). In addition, two subcellular localization datasets generated by proteomic studies of S. cerevisiae indicated that ScDGAT2 localizes in ER and lipid droplets (Huh et al., 2003; Natter et al., 2005). Moreover, recombinant expression of ScDGAT2 in a yeast strain devoid of TAG biosynthesis indicated that ScDGAT2 localizes in the microsomal fraction as an integral membrane protein (unpublished). Due to the presence of conserved transmembrane domains, it is expected that yeast DGAT2 localizes in the ER. The mechanisms involved in its movement from the ER to lipid droplets, however, are not yet determined. S. cerevisiae