The Metabolic Pathway Engineering Handbook: Fundamentals

THE METABOLIC PATHWAY ENGINEERING HANDBOOK Fundamentals

The Metabolic Pathway Engineering Handbook, 1st Edition The Metabolic Pathway Engineering Handbook: Fundamentals The Metabolic Pathway Engineering Handbook: Tools and Applications

THE METABOLIC PATHWAY ENGINEERING HANDBOOK Fundamentals

Edited by

Christina D. Smolke

CRC Press Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 © 2010 by Taylor & Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group, an Informa business No claim to original U.S. Government works Printed in the United States of America on acid-free paper 10 9 8 7 6 5 4 3 2 1 International Standard Book Number-13: 978-1-4398-0296-0 (Hardcover) This book contains information obtained from authentic and highly regarded sources. Reasonable efforts have been made to publish reliable data and information, but the author and publisher cannot assume responsibility for the validity of all materials or the consequences of their use. The authors and publishers have attempted to trace the copyright holders of all material reproduced in this publication and apologize to copyright holders if permission to publish in this form has not been obtained. If any copyright material has not been acknowledged please write and let us know so we may rectify in any future reprint. Except as permitted under U.S. Copyright Law, no part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, please access www.copyright.com (http:// www.copyright.com/) or contact the Copyright Clearance Center, Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a not-for-profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe. Library of Congress Cataloging-in-Publication Data The metabolic pathway engineering handbook : fundamentals / editor, Christina D. Smolke. p. ; cm. Includes bibliographical references and index. ISBN 978-1-4398-0296-0 (hardcover : alk. paper) 1. Genetic engineering--Handbooks, manuals, etc. 2. Biosynthesis--Handbooks, manuals, etc. I. Smolke, Christina D. II. Title. [DNLM: 1. Genetic Engineering--methods. 2. Metabolic Networks and Pathways. 3. Biological Products--metabolism. 4. Biotechnology--methods. 5. Models, Biological. QU 450 M5871 2010] TP248.6.M478 2010 660.6’5--dc22 Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com and the CRC Press Web site at http://www.crcpress.com

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Contents Introduction ............................................................................................................................... ix Editor . ......................................................................................................................................... xv Contributors . .......................................................................................................................... xvii

Section I  Cellular Metabolism Andy Ekins and Vincent J.J. Martin

1

Solute Transport Processes in the Cell . ................................................................... 1-1



2

Catabolism and Metabolic Fueling Processes ........................................................ 2-1



3

Biosynthesis of Cellular Building Blocks: The Prerequisites of Life . .............. 3-1



4 Polymerization of Building Blocks to Macromolecules:

Adelfo Escalante, Alfredo Martínez, Manuel Rivera, and Guillermo Gosset Olubolaji Akinterinwa and Patrick C. Cirino

Zachary L. Fowler, Effendi Leonard, and Mattheos Koffas

Polyhydroxyalkanoates as an Example .................................................................... 4-1 Si Jae Park, Soon Ho Hong, and Sang Yup Lee



5 Rare Metabolic Conversions—Harvesting Diversity

through Nature ............................................................................................................... 5-1 Manuel Ferrer and Peter N. Golyshin

Section II  Balances and Reaction Models Walter M. van Gulik

6

Growth Nutrients and Diversity ................................................................................ 6-1



7

Mass Balances, Rates, and Experiments .................................................................. 7-1



8

Data Reconciliation and Error Detection . .............................................................. 8-1



9

Black Box Models for Growth and Product Formation ....................................... 9-1

Joseph J. Heijnen Joseph J. Heijnen

Peter J.T. Verheijen Joseph J. Heijnen

v

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10

Contents

Metabolic Models for Growth and Product Formation ..................................... 10-1

Walter M. van Gulik

11 A Thermodynamic Description of Microbial Growth and

Product Formation . ..................................................................................................... 11-1

Joseph J. Heijnen

Section III Bacterial Transcriptional Regulation of Metabolism James C. Liao

12 13 14

Transcribing Metabolism Genes: Lessons from a Feral Promoter ................. 12-1 Alan J. Wolfe

Regulation of Secondary Metabolism in Bacteria ............................................... 13-1

Wenjun Zhang, Joshua P. Ferreira, and Yi Tang

A Synthetic Approach to Transcriptional Regulatory Engineering .. ............. 14-1

Wilson W. Wong and James C. Liao

Section IV  Modeling Tools for Metabolic Engineering Costas D. Maranas

15

Metabolic Flux Analysis ............................................................................................. 15-1

16

Metabolic Control Analysis . ..................................................................................... 16-1

17 18 19 20

Maria I. Klapa

Joseph J. Heijnen

Structure and Flux Analysis of Metabolic Networks ......................................... 17-1

Kiran Raosaheb Patil, Prashant Madhusudan Bapat, and Jens Nielsen

Constraint-Based Genome-Scale Models of Cellular Metabolism ................. 18-1

Radhakrishnan Mahadevan

Multiscale Modeling of Metabolic Regulation . .................................................. 19-1 C.A. Leclerc and Jeffrey D. Varner

Validation of Metabolic Models ............................................................................... 20-1

Sang Yup Lee, Hyohak Song, Tae Yong Kim, and Sung Bum Sohn

Section V Developing Appropriate Hosts for Metabolic Engineering Jens Nielsen

21 Escherichia coli as a Well-Developed Host for

Metabolic Engineering . .............................................................................................. 21-1

22

Eva Nordberg Karlsson, Louise Johansson, Olle Holst, and Gunnar Lidén

Metabolic Engineering in Yeast ............................................................................... 22-1 Maurizio Bettiga, Marie F. Gorwa-Grauslund, and Bärbel Hahn-Hägerdal

Contents

vii

23

Metabolic Engineering of Bacillus subtilis ............................................................ 23-1

24

Metabolic Engineering of Streptomyces .................................................................24-1

25 26

John Perkins, Markus Wyss, Hans-Peter Hohmann, and Uwe Sauer Irina Borodina, Anna Eliasson, and Jens Nielsen

Metabolic Engineering of Filamentous Fungi . .................................................... 25-1

Mikael Rørdam Andersen, Kanchana Rucksomtawin, Gerald Hofmann, and Jens Nielsen

Metabolic Engineering of Mammalian Cells . ......................................................26-1

Lake-Ee Quek and Lars Keld Nielsen

Index ........................................................................................................................................... I-1

Introduction

Progression of Biological Synthesis Methods toward Commercial Relevance The advent of recombinant DNA in the 1970s brought transformative technologies for the synthesis and manipulation of artificial genetic material. The ability to amplify, cut, and piece together fragments of DNA outside of a cell and to get (or transform) that DNA into a cell of interest resulted in a set of molecular cloning tools that enabled the field of genetic engineering. In genetic engineering, foreign DNA that encodes for new or altered functions or traits is inserted into an organism of interest. Many early applications of recombinant DNA technology focused on heterologous protein production in microbial hosts. The first medicine made through recombinant DNA technology that was approved by the United States Federal Drug Administration was the synthesis of synthetic “human” insulin in Escherichia coli. This was an important early application of recombinant DNA technology, as the success of producing a safe and effective synthetic hormone in a bacteria led to the widespread acceptance of the technology and significant resources and funding to be directed to its support and advancement. As the technologies in support of synthesizing and manipulating artificial DNA matured and advanced, so did the applications to which they were applied. The early successful applications of recombinant DNA technology resulted in alternative routes to the synthesis of medicines, such as insulin, human growth factor, and erythropoietin, vaccines, and even genetically modified organisms, including crops that exhibit more desirable traits. Technologies were developed for the manipulation of artificial DNA in both prokaryotic and eukaryotic host organisms, including mammalian and plant cells. In addition, inspired by the diversity of natural products, chemicals, and materials synthesized by biological systems that are observed in the natural world, researchers began to look beyond applications that were limited to the synthesis of a single heterologous protein product in a cellular host to more complicated engineering feats. In particular, these new applications focused on the manipulation of sets or combinations of proteins, or enzymes, that acted in conjunction in a cell, within metabolic pathways, to convert energy and precursor chemicals into desired natural and non-natural products. The production of chemicals, materials, and energy through biology presents an alternative to traditional chemical synthesis routes. While the development of chemical synthesis methods for the production of valuable chemicals and small molecule pharmaceuticals is a more mature field and has demonstrated significant successes, many chemicals remain difficult to be synthesized through such strategies, particularly those with many chiral centers. Biological catalysts, or enzymes, have demonstrated remarkable adeptness at the synthesis of very complex molecules. In addition, cellular biosynthesis strategies offer several advantages over traditional chemical synthesis strategies in that the former is often conducted under less harsh conditions, thereby enabling “green” synthesis strategies that are associated with the production of fewer toxic by-products. In addition, cellular biosynthesis ix

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Introduction

takes advantage of the cell’s natural ability to replenish enzymes and cofactors and to provide precursors from often inexpensive and renewable starting materials. Such advantages are particularly compelling in light of the global challenges we face today in energy, the environment, and sustainability. However, new challenges are presented when manipulating the metabolic pathways in cellular hosts that link energy sources and starting materials to products of commercial interest. The unique challenges faced in engineering metabolic pathways, when compared to the early genetic engineering applications of heterologous protein production, require the development of new enabling technologies, spanning experimental and analytical techniques and computational tools.

The Field of Metabolic Engineering Metabolic engineering is a field that includes the construction, redirection, and manipulation of cellular metabolism through the alteration of endogenous and/or heterologous enzyme activities and levels to achieve the biosynthesis or biocatalysis of desired compounds. Researchers in metabolic engineering often view the biological system as a chemical factory that is converting starting materials to different value-added products. Because the yield or productivity of the process is linked to its commercial viability, the ability to precisely regulate the flow of energy and materials through different cellular pathways becomes critical to the optimization of the overall process, drawing parallels to the more traditional engineering discipline of chemical process design. The basic tenet of metabolic engineering, the use of biology as a technology for the conversion of energy, chemicals, and materials to value-added products, has a long history. Early applications can be cited, even prior to the development of recombinant DNA technology, in the food and beverage industry where more traditional methods of strain development based on evolution, mating, and selection strategies were used to develop more desired production hosts for particular applications. However, recombinant DNA technology enabled the capability to introduce new enzymatic activities and pathways into production hosts allowing access to different energy resources and starting materials and to the production of different chemicals and materials. Such technologies support the forward design of more complex synthetic pathways in host organisms or the targeted manipulation of endogenous pathways, enabling more directed manipulation of the cellular host. Current metabolic engineering efforts are focused on the synthesis of products such as chemical commodities, small molecule drugs, and alternative energy sources including biofuels. In addition, significant effort is also directed to the engineering of host metabolisms to utilize renewable, low cost energy resources. Many of the challenges faced in metabolic engineering are related to the engineering of energy and material flow within complex systems. More specifically, metabolic pathways make up complex interconnected networks in cells, which can rarely be manipulated in isolation of the rest of the network. Highlighting the interconnections between cellular metabolites is the fact that all metabolites are made from a set of 12 common precursors. In addition, the flow of metabolites through a network of enzymes, and in the background of other cellular enzymes that may exhibit activity on these metabolites, is often controlled through layered processes that act at different time scales, implement dynamic feedback control, and utilize localization and transport. Metabolic engineering requires a breadth of skill sets to tackle different points of system design and as a result has developed into a very interdisciplinary field. Researchers with expertise spanning a variety of disciplines, including chemical engineering, biological engineering, environmental engineering, biochemistry, molecular biology, cell biology, bioinformatics, and control theory, are working in different areas of metabolic engineering. However, as an academic endeavor, metabolic engineering has remained an interdisciplinary research discipline with courses covering aspects of the field depending on the expertise of the department in which it is taught. As it has matured, metabolic engineering has gained greater industrial significance. Initial industrial interest was directed to the synthesis of chemical commodities in microorganisms largely at groups within larger chemical companies. However, many smaller startup companies have developed in recent years that are focused on the synthesis of specialty chemicals such as pharmaceuticals and biofuels, on

Introduction

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the development of computational and modeling programs to direct metabolic engineering efforts, and on the discovery and development of new enzyme activities in support of engineering new synthetic pathways into host organisms. The intersection of metabolic engineering, with other emerging areas of systems and synthetic biology, presents exciting opportunities to develop solutions to many of the global challenges we face in energy, the environment, health and medicine, resources, and sustainability, and will likely continue to fuel a significant sector of the biotechnology industry in future years.

An Overview of The Metabolic Pathway Engineering Handbook The purpose of The Metabolic Pathway Engineering Handbook is to provide a thorough overview of the field of metabolic engineering. Each section provides an overview of different aspects of a particular topic that is a central component of the field by experts in that area. Sections are introduced by section editors to provide a perspective on the topic and a description of how the chapters in that section link together to form an integrated overview of that particular topic. The sections are split into two books, where the content of the first book focuses on “fundamentals” or basic principles of metabolic engineering and the second book focuses on “tools and applications” in metabolic engineering. Due to its organization, the handbook can be used as a reference book and read for individual sections or chapters, or it can be used as a book for advanced courses in metabolic engineering. Section I in The Metabolic Pathway Engineering Handbook: Fundamentals provides an overview of the basic processes that support cellular metabolism. The boundary of a cell is defined by its cellular membrane, which acts to separate cellular constituents from the environment. Metabolism begins with systems that allow the import of nutrients and starting materials across the cellular membrane and efforts to engineer transport systems for particular chemicals have been important strategies in enabling cells to convert those chemicals to desired products. Once inside the cell, nutrients are broken down into common precursors for metabolic syntheses, which provide the energy and reducing power necessary for cell survival. In addition, precursors are channeled into the synthesis of important building blocks that the cell then utilizes to build larger macromolecules, including lipids, nucleic acids, and proteins. An understanding of the central metabolic pathways and the general flow of metabolism through a small number of common precursors and carriers is critical to being able to effectively link new synthetic nutrient or product pathways to endogenous metabolisms. Finally, the wealth of untapped diversity in nature, particularly in the microbial biosphere, provides significant opportunities in harvesting new enzymatic activities from nature that can be applied to the production of new chemicals and materials in engineered hosts. Section II provides an overview of mass balances and reaction models applied to predicting product formation and microbial growth in fermentation processes. Various models have been proposed and utilized in the field that exhibit varying levels of detail to provide predictions of product yield and cell growth. Conversion rates are calculated from mass balances and rate equations that take into account the basic nutrients and constituents of cellular systems. Different models, such as those based on thermodynamic or metabolic network constraints, can be utilized to predict product yield and cell growth in fermentation processes. Different models may be more or less appropriate based on the specifics of the fermentation. The application of such models to experimental systems can allow minimization of error in detection strategies resulting in optimized control schemes for fermentations based on such experimental measurements. Section III provides an overview of transcriptional regulation of metabolic pathways in bacterial systems. Bacterial cells use a variety of mechanisms to regulate the transcription of enzymes involved in primary and secondary metabolisms. Transcriptional regulatory strategies exist that regulate a small set of genes in response to specific environmental chemicals, such as operon-specific regulation and two-component systems. However, other strategies exist that regulate larger sets of genes in response to significant environmental changes such as heat shock or nitrogen starvation, through sigma factors and global transcriptional factors. An understanding of the strategies used to regulate the expression of

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Introduction

enzymes in a cellular host is critical in metabolic engineering to developing effective strategies to alter the expression of endogenous enzymes and to design synthetic systems that exhibit more sophisticated regulatory schemes to balance and coordinate the expression of multiple enzymes to ultimately optimize flux through desired pathways. Section IV is an overview of modeling tools that have been developed for metabolic engineering applications. Earlier modeling and computation efforts that resulted in tools for metabolic flux analysis (MFA) and metabolic control analysis (MCA) have been very powerful for the elucidation of fluxes and control strategies in metabolic networks given partial sets of data. Computation tools based on network and graph concepts have enabled structure and flux analyses that provide optimization tools for metabolic engineering. In addition, metabolic network reconstruction and modeling efforts have resulted in genome-scale models of cellular metabolism for specific organisms based on sets of constraints that enable prediction of flux distributions under different conditions. Whereas multi-scale modeling tools are extending current predictive capabilities by integrating stoichiometry, kinetics, and regulatory and control responses in metabolic networks, such multi-scale tools can be utilized by metabolic engineers to predict the dynamic metabolic response. Section V provides an overview of common cellular hosts that are used in metabolic engineering applications. In particular, the bacterial hosts Escherichia coli, Bacillus subtilis, and Streptomyces have been utilized in various metabolic engineering applications, with E. coli being the most well-developed and utilized host largely due to the genetic tools available for manipulating pathways in this host organism. In addition, two lower eukaryotic hosts, yeast and filamentous fungi, have been utilized in various metabolic engineering applications for the production of natural products or for pathway enzymes that are more readily expressed in functional forms in eukaryotic organisms. Finally, much effort has also been put toward the development of mammalian cell culture hosts for the production of metabolites and products that are more readily produced in mammalian cells. Each host may present advantages and disadvantages in the synthesis of a desired chemical based on the genetic tools available for manipulating pathways and the endogenous metabolism and processing pathways present in that organism, such that the selection of a suitable host is driven largely by the properties of the pathway of interest. Section I in The Metabolic Pathway Engineering Handbook: Tools and Applications provides an overview of the evolutionary tools widely in use in the engineering of metabolic enzymes and networks. Evolutionary strategies have been traditionally used in metabolic engineering to select for desired phenotypes in host organisms. As biological organisms naturally undergo processes of evolution and selection, design strategies that integrate evolutionary engineering objectives with metabolic engineering objectives may result in a more robustly performing engineered cellular system. Directed evolution is a laboratory tool that is used to mimic the evolutionary process in a test tube, by generating diversity in cellular components and then screening or selecting through this diversity for optimized component properties. Various experimental strategies have been utilized for generating and screening through component diversity. In addition, computational tools have been developed that optimize the design of laboratory evolution strategies. These experimental and computational tools have been applied to the directed evolution of enzymes, regulatory systems, pathways, and whole genomes for the optimization of flux through targeted metabolic pathways. Section II provides an overview of gene expression tools that have been utilized in metabolic engineering applications. Various tools have been developed that regulate DNA copy number and enable chromosomal engineering in host organisms. In addition, a variety of other genetic tools have been developed that precisely regulate gene expression levels through post-transcriptional and translational mechanisms. Still other tools have been developed that regulate the activity of enzymes through posttranslational engineering strategies. The application of the tools described in this section is critical to balancing the expression of multiple enzymes, such that individual conversion steps do not limit product yield, toxic intermediates do not accumulate, and cellular resources and energy are efficiently utilized by the host cell. Several examples exist of engineered systems that have utilized such genetic tools for the optimization of flux through metabolic pathways.

Introduction

xiii

Section III provides an overview of emerging technologies and their application to metabolic engineering. Genome-wide technologies that allow global profiling of cellular transcripts, proteins, metabolites, and phenotypes are critical for efficient troubleshooting and debugging of engineered systems. Bioinformatics tools that allow for management and analysis of the vast amounts of data collected from these techniques are also critical. As these technologies mature and become more available, their implementation as standard techniques in metabolic engineering will improve our understanding of the engineered system response and result in efficient troubleshooting and optimization strategies. Section IV provides an overview of key future prospects in metabolic engineering. The integration of new computational tools, such as genome-scale models, and new technologies for analyzing and understanding complex systems, such as systems biology, with metabolic engineering are rapidly advancing the success with which metabolic networks can be forward engineered. In addition, alternative strategies to cellular biosynthesis that remove complications associated with engineering living, evolving systems, such as cell-free synthesis systems, have demonstrated impressive successes. Finally, the modeling and optimization of engineered metabolic pathways in silico, prior to construction and characterization, will significantly transform the field of metabolic engineering and integrate advances in computational modeling, systems biology, and engineering design. Section V provides an overview of common tools that are utilized to determine flux through metabolic pathways. Various types of isotope flux labeling strategies have been widely used to monitor flux through metabolic pathways, where the data from such experiments are typically integrated into the modeling tools described in Section IV. In addition, various analytical strategies are utilized to profile cellular metabolites, where current and future efforts have been focused on developing strategies to profile and quantify global metabolite levels. Section VI provides an overview of various metabolic engineering application areas. One broad application area is focused on the engineering and regulation of the energy state, cofactor supply, and redox balance of cellular hosts. This is a challenge that affects most if not all metabolic engineering applications, where the introduction of new pathways or the manipulation of endogenous pathways can result in imbalances in cellular pathways and stress responses. Metabolic engineering applications are generally directed toward the synthesis of commercially relevant molecules including specialty or commodity chemicals, small molecule drugs, or alternative energy sources. Each of these application areas of metabolic engineering presents distinct challenges that must be addressed in the process design based on chemical and pathway complexity, market cost of the product, volume demand of the product, end use of the product, and purity requirements.

Metabolic Engineering: Looking toward the Future Metabolic engineering as a field has evolved significantly over the past 10 to 15 years in large part due to the scientific and technological advances made during this time frame in support of this application area. The future prospects of metabolic engineering are extremely exciting, and as other supporting scientific and engineering fields mature it is likely to see transformative advances that direct it further toward an engineering discipline. There are several key supporting fields that will aid in directing this transformation. First, enzyme engineering and enzyme discovery will be critical to expanding the diversity of natural and non-natural products that can be produced in engineered organisms. Much of the living world has not been cultured and characterized. Even in those organisms that have been cultured, we do not have genome sequence information, have not mapped functions to many of the sequenced genes, or have not characterized many of the enzyme activities in these organisms. For example, many pathways in plants responsible for the synthesis of diverse pharmacologically relevant molecules have not been elucidated, although many of these activities and their corresponding genes are currently present in large expressed sequence tag (EST) libraries. Because we cannot forward design enzymes to exhibit specific catalytic

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activities, the existing limitations in characterized enzyme activities severely limit the pathways that we can reconstruct in organisms. In addition, programs that will allow us to predict and design enzyme function from sequence will be critically enabling for the design of new activities that have not been recovered from natural systems. Second, because metabolic engineering is largely a systems engineering challenge, continued advances in systems biology will provide important insights into the function of biological systems that will inform engineering design and strategies directed at manipulating metabolic pathways. Many analytic techniques in support of systems biology, including strategies that allow global profiling of transcript, protein, and metabolite levels, are providing vast amounts of information regarding levels of cellular constituents under different conditions. In addition, computational tools are being developed to process the vast amounts of data coming from these techniques. Newer and future efforts in systems biology must focus on taking the information coming from these techniques and abstracting from it the organizing principles governing cellular metabolism and regulation. An understanding of how cells generally layer metabolic pathways with different regulatory strategies will allow engineers to design more robustly performing synthetic pathways that are better integrated with the endogenous metabolic pathways. In addition, such understanding will allow better identification of manipulation points in endogenous networks to alter flux through pathways. Third, the integration of information theory and control theory with systems biology and metabolic engineering will likely have a significant impact on our understanding of biological systems. Such tools will enable a deeper understanding of architectures and properties of complex networks that support robustness, evolvability, and fragility of the system, providing a conceptual framework to systems biology. In addition, such tools will allow researchers to more quantitatively examine models of control schemes around metabolic pathways to better elucidate the design principles around regulating flux through metabolic pathways. Such tools can also be used to examine synthetic network and control scheme designs and guide the more effective design of engineered systems. Finally, metabolic engineering is seeing a transformation with the emerging field of synthetic biology. Synthetic biology is the design, construction, and characterization of biological systems using engineering design principles. To support a framework for engineering biology, synthetic biology is rooted in foundational technologies that enable the construction of more complex, heterologous networks in living systems. With advances in DNA sequencing and synthesis it is becoming common practice to synthesize entire genes and pathways from scratch, no longer limiting researchers to the physical DNA that they obtain from natural organisms. In addition, abstraction frameworks have been proposed to enable rapid assembling and reassembling of basic biological components (or parts) into larger networks (or devices) and systems, supporting the rapid prototyping and troubleshooting and reliable construction of complex metabolic pathways in cellular hosts (or chassis). An example of a synthetic biology approach to the rapid prototyping of a metabolic pathway in Escherichia coli was recently described (http://parts. mit.edu/wiki/index.php/MIT_2006). There are also efforts directed to the engineering of specific chassis, or cellular hosts, optimized for metabolic engineering applications. Finally, enabling genetically encoded technologies are being developed for use in precise and quantitative manipulation of pathway components such as enzymes.

Christina D. Smolke Editor-in-Chief

Editor

Christina Smolke is an assistant professor in the Department of Bioengineering at Stanford University. She graduated with a BS in chemical engineering with a minor in biology from the University of Southern California in 1997. She conducted her graduate training as a National Science Foundation Fellow in the Chemical Engineering Department at the University of California at Berkeley and earned her PhD in 2001. Christina conducted her postdoctoral training as a National Institutes of Health Fellow in cell biology at UC Berkeley. She started her independent research program as an assistant professor in the Division of Chemistry and Chemical Engineering at the California Institute of Technology from 2003– 2008. She has pioneered a research program in developing foundational technologies for the design and construction of engineered ligand-responsive RNA-based regulatory molecules, their integration into molecular computation and signal integration strategies, and their reliable implementation into diverse cellular engineering applications. These technologies are resulting in scaleable platforms for the construction of molecular tools that work across many cellular systems and allow regulation of targeted gene expression levels in response to diverse endogenous or exogenous molecular ligands. Her research is rapidly advancing current capabilities of noninvasive detection of cellular state and programming cellular function. In particular, her laboratory is examining the application of these tools to the optimization of metabolic pathway engineering strategies in organisms such as yeast. Dr. Smolke’s innovative research program has recently been recognized with the receipt of a National Science Foundation CAREER Award, a Beckman Young Investigator Award, an Alfred P. Sloan Research Fellowship, and the listing of Dr. Smolke as one of Technology Review’s Top 100 Young Innovators in the World. She is also a member and adjunct faculty of the Comprehensive Cancer Center’s Cancer Immunotherapeutics Program at the City of Hope, where she has several translationally oriented collaborative projects exploring the clinical applications of these technologies. She is the inventor of over nine patents and serves on the Scientific Advisory Board of Codon Devices. Dr. Smolke is currently serving as the President of the Institute of Biological Engineering. She is a member of AIChE, ACS, the RNA Society, and IBE.

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Contributors

Olubolaji Akinterinwa

Andy Ekins

Peter N. Golyshin

Mikael Rørdam Andersen

Anna Eliasson

Marie F. GorwaGrauslund

Department of Chemical Engineering Pennsylvania State University University Park, Pennsylvania

Department of Biology Centre for Structural and Functional Genomics Concordia University Montreal, Quebec, Canada

Center for Microbial Biotechnology BioCentrum-DTU Technical University of Denmark Lyngby, Denmark

Center for Microbial Biotechnology BioCentrum-DTU Technical University of Denmark Lyngby, Denmark

Prashant Madhusudan Bapat

Adelfo Escalante

Maurizio Bettiga

Joshua P. Ferreira

Center for Microbial Biotechnology BioCentrum-DTU Technical University of Denmark Lyngby, Denmark

Department of Applied Microbiology Lund University Lund, Sweden

Irina Borodina

Center for Microbial Biotechnology BioCentrum-DTU Technical University of Denmark Lyngby, Denmark

Patrick C. Cirino

Department of Chemical Engineering Pennsylvania State University University Park, Pennsylvania

Cellular Engineering Biocatalysis Department Biotechnology Institute National Autonomous University of Mexico Cuernavaca, Mexico Department of Chemical and Biomolecular Engineering University of California Los Angeles, California

Manuel Ferrer

Department of Biocatalysis Institute of Catalysis Consejo Superior de Investigaciones Científicas Madrid, Spain

Zachary L. Fowler

Department of Chemical and Biological Engineering State University of New York at Buffalo Buffalo, New York

Department of Environmental Microbiology HZI-Helmholtz Centre for Infection Research Braunschweig, Germany

Department of Applied Microbiology Lund University Lund, Sweden

Guillermo Gosset

Cellular Engineering Biocatalysis Department Biotechnology Institute National Autonomous University of Mexico Cuernavaca, Mexico

Bärbel Hahn-Hägerdal Department of Applied Microbiology Lund University Lund, Sweden

Joseph J. Heijnen

Bioprocess Technology Group Department of Biotechnology Delft University of Technology Delft, the Netherlands

Gerald Hofmann

Center for Microbial Biotechnology, BioCentrum-DTU Technical University of Denmark Lyngby, Denmark

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Hans-Peter Hohmann

DSM Nutritional Products Ltd Basel, Switzerland

Olle Holst

Department of Biotechnology Lund University Lund, Sweden

Contributors

C.A. Leclerc

Department of Chemical Engineering McGill University Montreal, Quebec, Canada

Department of Chemical Engineering and Bioengineering University of Ulsan Ulsan, Republic of Korea

Louise Johansson

Effendi Leonard

Department of Chemical Engineering Lund University Lund, Sweden

Department of Chemical and Biological Engineering State University of New York at Buffalo Buffalo, New York

Eva Nordberg Karlsson Department of Biotechnology Lund University Lund, Sweden

Tae Yong Kim

Department of Chemical and Biomolecular Engineering Center for Systems and Synthetic Biotechnology Institute for the BioCentury Korea Advanced Institute of Science and Technology Daejeon, Korea

Maria I. Klapa

James C. Liao

Chemical and Biomolecular Engineering Department University of California Los Angeles, California

Gunnar Lidén

Department of Chemical Engineering Lund University Lund, Sweden

Radhakrishnan Mahadevan

Department of Chemical and Biomolecular Engineering Institute of Chemical Engineering and High-Temperature Chemical Processes Foundation for Research and Technology-Hellas Patras, Greece

Department of Chemical Engineering and Applied Chemistry Institute of Biomaterials and Biomedical Engineering University of Toronto Toronto, Ontario, Canada

Mattheos Koffas

Costas D. Maranas

Department of Chemical and Biological Engineering State University of New York at Buffalo Buffalo, New York

Department of Biology Centre for Structural and Functional Genomics Concordia University Montreal, Quebec, Canada

Sang Yup Lee

Department of Chemical and Biomolecular Engineering Center for Systems and Synthetic Biotechnology Institute for the BioCentury Korea Advanced Institute of Science and Technology Daejeon, Korea

Soon Ho Hong

Vincent J.J. Martin

Department of Chemical Engineering Pennsylvania State University Fenske Laboratory University Park, Pennsylvania

Alfredo Martínez

Cellular Engineering Biocatalysis Department Biotechnology Institute National Autonomous University of Mexico Cuernavaca, Mexico

Jens Nielsen

Systems Biology Department of Chemical and Biological Engineering Chalmers University of Technology Gothenburg, Sweden and Center for Microbial Biotechnology BioCentrum-DTU Technical University of Denmark Lyngby, Denmark

Lars Keld Nielsen

Australian Institute for Bioengineering and Nanotechnology The University of Queensland Brisbane, Australia

Si Jae Park

Corporate R&D LG Chem, Ltd. Daejeon, Republic of Korea

Kiran Raosaheb Patil

Center for Microbial Biotechnology BioCentrum-DTU Technical University of Denmark Lyngby, Denmark

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Contributors

John Perkins

DSM Nutritional Products Ltd Basel, Switzerland

Lake-Ee Quek

Australian Institute for Bioengineering and Nanotechnology The University of Queensland Brisbane, Australia

Manuel Rivera

Cellular Engineering Biocatalysis Department Biotechnology Institute National Autonomous University of Mexico Cuernavaca, Mexico

Kanchana Rueksomtawin

Center for Microbial Biotechnology BioCentrum-DTU Technical University of Denmark Lyngby, Denmark

Uwe Sauer

Institute for Molecular Systems Biology ETH Zürich Zürich, Switzerland

Christina D. Smolke

Jeffrey D. Varner

Seung Bum Sohn

Peter J.T. Verheijen

Division of Chemistry and Chemical Engineering California Institute of Technology Pasadena, California

Department of Chemical and Biomolecular Engineering Center for Systems and Synthetic Biotechnology Institute for the BioCentury Korea Advanced Institute of Science and Technology Daejeon, Korea

Hyohak Song

Department of Chemical and Biomolecular Engineering Center for Systems and Synthetic Biotechnology Institute for the BioCentury Korea Advanced Institute of Science and Technology Daejeon, Korea

Department of Chemical and Biomolecular Engineering Cornell University Ithaca, New York

Department of Biotechnology Delft University of Technology Delft, the Netherlands

Alan J. Wolfe

Department of Microbiology and Immunology Loyola University at Chicago Stritch School of Medicine Maywood, Illinois

Wilson W. Wong

Chemical and Biomolecular Engineering Department University of California Los Angeles, California

Yi Tang

Department of Chemical and Biomolecular Engineering University of California Los Angeles, California

Walter M. van Gulik

Bioprocess Technology Group Department of Biotechnology Delft University of Technology Delft, the Netherlands

Markus Wyss

DSM Nutritional Products Ltd Basel, Switzerland

Wenjun Zhang

Department of Chemical and Biomolecular Engineering University of California Los Angeles, California

Cellular Metabolism

I

Andy Ekins and Vincent J.J. Martin Concordia University

1 Solute Transport Processes in the Cell  Adelfo Escalante, Alfredo Martínez, Manuel Rivera, and Guillermo Gosset......................................................................................1-1 Introduction  •  Structure and Function of the Bacterial Membrane  •  The Transporter Classification (TC) System

2 Catabolism and Metabolic Fueling Processes  Olubolaji Akinterinwa and Patrick C. Cirino...........................................................................................................................2-1 Introduction  •  Classification of Organisms  •  Thermodynamics of Fueling Processes  •  Products of Fueling Processes  •  Redox Potentials and Mobile Electron Carriers  •  Examples of Catabolic Processes in Different Organisms  •  Concluding Remarks

3 Biosynthesis of Cellular Building Blocks: The Prerequisites of Life  Zachary L. Fowler, Effendi Leonard, and Mattheos Koffas......................................3-1 Introduction  •  Amino Acid Biosynthesis  •  Nucleotides as Building Blocks  •  Synthesis of Carbohydrates for Building Cells  •  Cell Synthesis of Lipids

4 Polymerization of Building Blocks to Macromolecules: Polyhydroxyalkanoates as an Example  Si Jae Park, Soon Ho Hong, and Sang Yup Lee.................................................................................................................................4-1 Introduction  •  PHAs  •  PHA Synthases  •  Metabolic Engineering of Microorganisms for PHA Production  •  Conclusion

5 Rare Metabolic Conversions—Harvesting Diversity through Nature  Manuel Ferrer and Peter N. Golyshin........................................................5-1 Introduction  •  How Diverse Are Functional Groups?  •  Diversity of Enyzmes and Current Frontiers for Bioconversions  •  Main Chemical Conversions Mediated by Enzymes: Putative “Rare” Conversions  •  How Can New Catalytic Functions Be Achieved?  •  Recent Advances in Metagenomics: The Untapped Reservoir of Proteins from Unculturable Microbes

I-1

I-2

T

Cellular Metabolism

he field of metabolic engineering has advanced over time and will undoubtedly continue to do so, based on a solid scientific foundation and continued research and innovation. Of critical importance is a solid understanding of the metabolism of the cell which will ultimately be manipulated through various techniques to produce a desired end product, or alternatively, remove or breakdown an undesirable one. This section describes the well defined knowledge of how, in particular Escherichia coli, is able to transport a variety of nutrients and use such nutrients, through a variety of metabolic pathways, to derive energy and synthesize the wide spectrum of cellular components required for the maintenance of a cell. In some instances, the alteration of such metabolic pathways of a particular cell may lead to the production of a desired product, while in others the synthesis of the desired product may require the introduction of foreign genes, isolated and characterized from other organisms, in order to allow synthesis to proceed. Furthermore, with the advent of metagenomics, one can sift out genes allowing unique metabolic conversions which have yet to have been described in cultured microorganisms. While the organism of choice for many metabolic engineering studies is E. coli, all cells are, by definition, enveloped by a membrane which separates cellular components from the extracellular environment. Highly efficient transport and export systems have evolved to allow exchange across this barrier. A sound knowledge of the transport systems which import the nutrients required for cell growth and drive the metabolism of the cell is essential in order to ensure that the desired metabolic pathway receives the necessary precursors and energy required for the production of a selected compound. As the outer membrane of E. coli is only capable of allowing the passive diffusion of molecules with a molecular weight less than approximately 600 Da, it is of utmost importance to determine if the import of precursors, for instance, can cause a bottleneck in the synthesis of a particular compound. Additionally, the type of transport system present in the cell can have an impact on the carbon flux within a cell. As an example, modification of the E. coli phosphotransferase system has been a strategy successfully applied to metabolically engineered strains (Gosset, 2005). Manipulation of the transport systems can increase the diversity of nutrients imported while manipulation of the regulatory systems of the cell can allow the simultaneous import and use of multiple carbon sources, as is the case for carbon catabolite repression mutants for example (Dien et al., 2002). As nutrients are catabolized, precursors for metabolic syntheses are generated along with the energy and reducing power required to drive the synthesis of all the components required by the cell. Solid knowledge of the metabolic pathways within the cell allows one to ensure that the proper precursors, reducing power and energy are in ample supply in order to produce the molecule of interest. Furthermore, culture components and conditions can be altered to enhance the efficiency of a particular metabolic conversion. In some instances, one may wish to overproduce a compound in an organism that does not naturally produce said compound. One such example is the production of polyhydroxyalkanoates (PHAs) in E. coli, an organism that does not naturally produce PHAs. It is therefore necessary to express PHA synthase genes isolated from foreign organisms. Additionally, in order to increase production of the desired PHA, it is crucial to evaluate the metabolic flux of the host organism and perhaps amplify certain endogenous pathways to increase the availability of precursors and reducing power, without decreasing the overall health of the expressing organism. In the case of poly (3-hydroxybutyrate) [P(3HB)] production, it was found that over expression of glucose-6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase led to an increase in the NADPH/NADP+ ratio and a subsequent rise in the concentration of P(3HB), there was however a detrimental effect on the producing cells and the observed increase in P(3HB) production was due to a lower cell concentration (Lim et al., 2002). In another experiment, 2-D gel electrophoresis and metabolic flux analysis was performed on E. coli producing P(3HB) and it was revealed that there was an increase in certain glycolytic pathway enzymes. Subsequently, amplification of the glycolytic pathway enzymes led to increased production of acetyl-CoA, which could subsequently be used to increase yields of P(3HB) (Park and Lee, unpublished results).

Cellular Metabolism

I-3

The pursuit of rare conversions have, up to this point, focused on the ability of cultured organisms and their enzymes to perform such functions. The products of such rare conversions are invaluable to a variety of industries spanning the agricultural, pharmaceutical, food additive and bioremediation fields, as examples. While many advances have been made in the realm of protein engineering using techniques such as directed evolution, it is reasonable to assume that an even greater diversity exists within the genomes of unculturable microorganisms. The diversity of the cultured microbial world has led to the discovery of many rare conversions, there remains, however, a large pool of untapped genetic material within the many “unculturable” microorganisms that are currently estimated to represent close to 99% of the microbial world (Fütterer et al., 2004). With the knowledge that each sequenced microbial genome yields on average 30–50% of genes with unknown function (Bode and Müller, 2005) and the recent shotgun sequencing of DNA isolated from the Sargasso Sea revealed greater than 1.2 million genes of unknown function (Venter et al., 2004) it would appear reasonable to assume that there exists a vast pool of untapped genetic resources that can be applied to all realms of biotechnology.

References Bode, H.B., and Müller, R. The impact of bacterial genomics on natural product research. Angew. Chem. Int. Ed. Engl., 44, 6828, 2005. Dien, B.S., Nichols, N.N., and Bothast, R.J. Fermentation of sugar mixtures using Escherichia coli catabolite repression mutants engineered for production of L-lactic acid. J. Ind. Microbiol. Biotechnol., 29, 221, 2002. Fütterer, O., et al. Genome sequence of Picrophilus torridus and its implications for life around pH 0. Proc. Natl. Acad. Sci. USA., 101, 9091, 2004. Gosset, G. Improvement of Escherichia coli production strains by modification of the phosphoenolpyruvate:sugar phosphotransferase system. Microb. Cell. Fact., 4, 14, 2005. Lim, S.J. et al. Amplification of the NADPH-related genes zwf and gnd for the oddball biosynthesis of PHB in an E. coli transformant harboring a cloned phbCAB operon. J. Biosci. Bioeng. 93, 543, 2002. Venter, J.C. et al. Environmental genome shotgun sequencing of the Sargasso Sea. Science, 304, 66, 2004.

1 Solute Transport Processes in the Cell

Adelfo Escalante, Alfredo Martínez, Manuel Rivera, and Guillermo Gosset National Autonomous University of Mexico

1.1 1.2

Introduction ���������������������������������������������������������������������������������������1-1 Structure and Function of the Bacterial Membrane.....................1-2

1.3

The Transporter Classification (TC) System..................................1-8

Structure of the Cellular Membrane  •  Functions  •  Kinetics of Transport Processes Channels and Pores  •  Electrochemical Potential-Driven Transporters  •  Primary Active Transporters  •  Group Translocators  •  Transmembrane Electron Flow Systems

References ����������������������������������������������������������������������������������������������������1-19

1.1  Introduction The cell membrane constitutes a hydrophobic barrier that isolates the cytoplasm from the external medium. The entry and exit of most of the nutrients required for cell growth and the byproducts generated by metabolism are highly restricted by this cellular structure. However, to sustain high growth rates, microbes require a high rate of nutrient import. The presence of specialized transport proteins in the membrane allows the cell to circumvent the permeability restrictions imposed by this barrier. Analyses of microbial genomes have revealed that approximately 10% of the genes encode proteins involved in transport [1]. These transport systems participate in the import and export of different classes of molecules and also in other important cellular functions. They allow the entry of nutrients to sustain metabolism and ion species to maintain concentration gradients leading to membrane potential and energy generation. Transporters allow the excretion of metabolite by-products and other toxic substances, like drugs or certain metal ions. Transport systems also participate in the secretion of lipids, carbohydrates, and proteins into membrane(s) or the external medium. They enable the transfer of nucleic acids between organisms, contributing to microbial diversity. Finally, transporters participate in the uptake of different types of signaling molecules like alarmones and hormones, among others, thus allowing cellular communication [2]. Solute transport and metabolism are linked processes in the cell. Genetic organization in bacteria frequently reflects this functional coupling by the clustering of genes encoding both transport and metabolic activities in transcriptional units. This association is generally observed in operons encoding catabolic pathways for carbon sources [3]. Transport and regulatory systems participate in the process whereby the bacterial cell can select from a mixture of nutrients those that afford the highest growth rate [4]. In addition, the differential expression of genes encoding distinct transporters for a specific substrate allow the cell to select the transport mechanism according to the physiological state and environmental conditions [5]. Transport systems are potential targets for modification with the aim of microbial production strain improvement. Metabolic engineering efforts usually focus on modifying metabolic enzyme activities. 1-1

1-2

Cellular Metabolism

However, it can be envisioned that high performance production strains will also require the modification of other cellular functions, including transport. Modification of transport systems can result in the improvement of several cellular properties including: (a) increasing the range of carbon source utilization [6]; (b) increasing metabolic precursor availability for the synthesis of amino acids, shikimate pathway intermediates, TCA cycle intermediates, and fermentation products like ethanol [7–10]; (c) increasing the efficiency in sugar mixture utilization by partial disruption of catabolic repression [11]; and (d) controlling overflow metabolism, thus reducing acetate production [12].

1.2  Structure and Function of the Bacterial Membrane 1.2.1  Structure of the Cellular Membrane The cell membrane, also known as the cytoplasmic membrane, plasma membrane, or cell surface membrane, is a thin structure that surrounds the cell. It is the barrier that defines the boundaries of the cell, separating the cytoplasm from its environment. If the membrane is damaged, the integrity of the cell is altered and the cytoplasm leaks into the environment, causing cell death. The general structure of prokaryotic and eukaryotic cell membranes (and the outer membranes of Gram-negative bacteria) is a bilayer composed of phospholipids, which contain both hydrophobic (fatty acid) and hydrophilic (glycerol-phosphate) components. It can exist in many chemical forms as a consequence of a diversity of compounds attached to the glycerol backbone. As phospholipids aggregate in aqueous solution they spontaneously organize to form two parallel rows, known as a lipid bilayer. Phospholipid molecules align with the fatty acids pointing inward toward each other to form a hydrophobic environment, whereas the hydrophilic portions face both the external side and the internal or cytoplasmic side of the membrane. The bilayer structure represents the most stable arrangement of the lipid molecules in an aqueous environment. The whole structure of the plasma membrane is stabilized by hydrogen bonds and hydrophobic interactions. In addition, cations such as Mg2 + and Ca 2 + help to stabilize the membrane due to ionic interactions with the negative charges of the phospholipids. A model of the structures of the bacterial cell membranes of Gram-positive and Gram-negative bacteria is shown in Figure 1.1 [13–15]. An important amount of protein and other materials is partially or completely embedded in the membrane layer. A typical bacterial membrane contains up to 200 different kinds of proteins (approximately 75% of the mass of the membrane). Protein molecules in the membrane are arranged in a variety of ways. Some proteins are fully embedded in the membrane and are thus called integral or transmembrane proteins. They can be removed from the membrane only after disrupting the lipidic bilayer. Some of these proteins are channels that have a pore, through which substances enter and exit the cell. Other proteins, called peripheral, are easily removed from the membrane by mild treatments and are firmly associated with the inner or outer surface of the membrane. They may function as enzymes that catalyze chemical reactions, as scaffolds for support of cell components, and as mediators of changes in membrane shape during movement. Some peripheral membrane proteins contain a lipid tail on the amino terminus that anchors the protein to the membrane. These proteins are called lipoproteins and interact directly with integral proteins in important cellular processes such as energy metabolism. Many proteins and some of the lipids on the outer membrane of the plasma membrane have carbohydrates attached to them. These structures are known as glycoproteins and glycolipids, respectively. Both of these structures help to protect the cell and are involved in cell-to-cell interactions. Sterols and related molecules are present in eukaryotic membranes. They are rigid and planar molecules, whereas fatty acids are flexible; their presence stabilizes and makes membranes less flexible. Sterols are absent in prokaryotic cellular membranes, except for methanotrophs and mycoplasms. Polycyclic compounds known as hopanes (derivatives of pentacyclic triterpenoides) are widely distributed among bacteria, and it is proposed that they may play a role in maintaining membrane rigidity (Figure 1.2). One widely distributed hopane is the C30 hopanoid diploptene. Hopanes are not present in species of Archaea [16].

1-3

Solute Transport Processes in the Cell O-specific side chains

Gram-negative

Lipopolysaccahride

Outer membrane

Murein lipoprotein

Periplasmic space and cell wall

Murein Phospholipids

Cytoplasmic membrane

Peripheral proteins

Transmembrane proteins

Gram-positive Cytoplasmic membrane

Figure 1.1  Cell membranes of Gram-positive and Gram-negative bacteria. Schematic representation of the inner and outer membrane lipid bilayers of Gram-negative bacteria (upper panel) and Gram-positives (lower panel). Several structures associated to cell membranes such as porins, integral or transmembrane and peripheral proteins, and cell wall components are shown. (a)

CH3 H3C

C

CH3 H2C

H2C

H2C CH CH3

H CH3

HO OH

(b)

CH3

CH3

CH3

OH

OH

OH

CH3

CH3

CH3

Figure 1.2  Structure of membrane sterols and hopanoids. (a) Structure of the cholesterol molecule, a typical sterol present in cell membranes of eukaryotic cells, methanotrophic bacteria, and mycoplasmas. (b) Structure of a hopane, a polyterpenoid present in prokaryotic cell membranes.

1-4

Cellular Metabolism

Membranes have a viscosity similar to that of light-grade oil. Experimental evidence has demonstrated that at temperatures that permit growth, membrane molecules are not static but move quite freely within the membrane surface. Individual lipid molecules are also generally free to exchange places with another lipid in the membrane, resembling a two dimensional fluid. It is proposed that this movement is most probably associated to the functions of the plasma membrane. These dynamics of phospholipids and proteins are known as the fluid mosaic model [17]. However, it is also proposed that some membrane regions have considerable order, because some lipids molecules are not free due to their relationship with specific membrane proteins and some other components [18]. The phospholipids of the cell membrane from bacteria contain ester linkages bonding the fatty acids to glycerol whereas in Archaea the membrane lacks fatty acids (Figure 1.3). Instead, their side chains are composed of repeating units of the five carbon hydrocarbon isoprene that is linked to glycerol by an ether bond; however, the overall architecture of the cytoplasmic membrane of Archaea, forming an inner and outer hydrophilic surfaces with a hydrophobic interior, is the same as in bacteria. Glycerol diethers and glycerol tetraethers are the major lipids present in membranes from Archaea. In the tetramer molecule, the phytanyl side chains (composed of four linked isoprenes) from each glycerol molecule are covalently bonded together (Figure 1.4), leading a lipid monolayer instead of a bilayer cytoplasmic membrane. This structure is widely distributed among hyperthermophilic Archaea helping to maintain the membrane architecture at high temperatures [19–20].

1.2.2  Functions The most important function of the cell membrane is to serve as a selective barrier through which material enters and exits the cell. The cytoplasm consists of an aqueous solution containing salts, sugars, amino acids, nucleotides, vitamins, coenzymes, proteins, and a variety of other soluble materials. The hydrophobic nature of the internal region of the plasma membrane constitutes a tight diffusion barrier with selective permeability, allowing certain molecules and ions to pass through and blocking passage to others. Some smaller molecules, such as water, oxygen, carbon dioxide, and some simple sugars, usually pass freely through the membrane by diffusion (Table 1.1). This is also the case for molecules that are dissolved easily in lipids (oxygen, carbon dioxide, and nonpolar organic molecules). In contrast, hydrophilic and small charged molecules such as the hydrogen ion (H+) do not pass through the membrane but instead must be specifically transported. Water is a molecule that freely crosses the membrane, because it is sufficiently small to pass through the phospholipid bilayer. However, water transport through the membrane can be accelerated

(a)

(b)

Ester

(c)

O H2C

O

C

Ether

R H2C

O

C

R

HC

O

C

R

H2C

O

O HC

O

C

R

O H2C

O

P O–

CH3

O O–

P

O–

H2C

C

C H

CH2

O–

Figure 1.3  Chemical diversity of lipidic bonds in cell membranes. (a) An ester linkage found in lipids of ­bacteria and eukaryotic cells. (b) An ether linkage of lipids of cell membrane of Archaea. (c) Structure of isoprene, the parent structure of the hydrophobic side chains present in Archaea.

1-5

Solute Transport Processes in the Cell (a) Glycerol diether

Phytanyl

Ether linkage

H

H H

Glycerol phosphate

C

O

CH2

CH2

H

C

O

CH2

H

C

O

R

Phytanyl

H

O

C

H

CH2

O

C

H

R

O

C

H

H

CH3 group

Lipid bilayer (b) Diglycerol tetraether

H H

H

O

C

H

H

C

O

CH2

CH2

O

C

H

H

C

O

CH2

CH2

O

C

H

H

C

O

H

H

Biphytanyl

H

Lipid monolayer

Figure 1.4  Structures of the Archaea cell membranes. (a) Schematic representation of bilayers of isoprenoids linked to glycerol by ether bonds. (b) Structure of the monolayers of the isoprenoid biphytanyl glycerol ether.

Table 1.1  Comparison of Diffusion-Controlled and Carrier Mediated Solute Fluxes across Bacterial Plasma Membranes Typical Transfer Rate [µmol min-1 (g Dry Mass) -1] Diffusion-Controlled at a Concentration Difference of Transported Solute Potassium ion (K ) Glutamate +

Glucose Isoleucine Phenylalanine Urea

10 µM

10 mM

Carrier Mediated (Vmax)

0.00002

0.02

100

<0.00005 0.001 0.0015 0.08 0.04

<0.05 1.0 1.5 8.0 40.0

50 30 1 5

25

by specific transport proteins called aquaporins. The movement of most hydrophilic solutes across plasma membranes depends on transporter molecules, which will be described in the following sections [21].

1.2.3  Kinetics of Transport Processes A solute transport system consists of integral membrane proteins that can be regarded as membranebound enzymes. However, instead of catalyzing the conversion of substrates to products, they mediate

1-6

Cellular Metabolism

the transfer of solutes between compartments separated by a membrane. Each transport system displays an affinity and specificity toward particular substrate(s). It is not uncommon to find in one organism more than one transporter for a solute, each having a different affinity and specificity, a situation analogous to that of isozymes in metabolism. In addition, there is considerable diversity with regard to energetic coupling mechanisms that drive active transport. Although transport processes are not fully understood, some models are helpful to understand basic molecular events [22,23]. The function of a transporter can be defined in three basics steps, analogous to those of enzyme activity: binding, translocation, and release of solute. The translocation step can involve a major conformational change of the transporter protein, thus defining either of two functional conformations. Diffusion plays an important role in solute transport across the lipid bilayer membrane. This process can occur either in the absence of or mediated by specific protein transporters. By measuring the transport rate of a solute it is possible to determine if the diffusion process is transporter mediated. In a transporter-independent process, there is a linear increase in the rate of diffusion with an increase in solute concentration, as shown in Figure 1.5. In contrast, in transporter mediated diffusion, a maximum value is reached. This response is explained considering that the transporter proteins become saturated once the solute substrate reaches a specific high concentration. The processes that govern the simple diffusion of an electrically neutral molecule were studied by Adolf Eugen Fick in the 1800s and is applied to the special case of a cell membrane. The measurement of the concentrations of a solute Sx outside and inside a cell allows us to predict if it is in equilibrium across the cell membrane or whether Sx would tend to passively move into the cell or out of it. As long as the movement of Sx is not coupled to the movement of another substance or to some biochemical reaction, the only factor determining the direction of the net transport is the difference in concentration. The ability to predict the movement of Sx is independent of any detailed knowledge of the actual transport pathway mediating its passive transport. When the concentration of an external solute Sx ([Sx]o) is greater than its internal concentration ([Sx]i), the net movement of Sx will be into the cell. The movement of Sx is described by its flux (Jsx), namely, the number of moles of Sx crossing a unit area of membrane (typically 1 cm2) per unit of time (s), i.e., moles/ (cm2∙s). High Sx solubility in the membrane lipids (the higher the lipid–water partition coefficient of Sx) will correspond to a higher flux through the membrane barrier. The flux of Sx will also be greater if Sx moves more readily once it is in the membrane (higher diffusion coefficient) and if the distance that it must traverse is short (membrane thickness). These three factors form part of the parameter called the permeability coefficient of Sx (Psx). Finally, the flux of Sx will be greater as the difference in [Sx] between the two sides of the membranes increases (gradient). All these concepts are integrated in the equation known as Fick´s law:

Rate of diffusion

Carrier mediated transport plus diffusion Carrier mediated transport

Diffusion

Substrate concentration (S)

Figure 1.5  Comparison of solute transport kinetics in the presence and absence of a transporter.

Solute Transport Processes in the Cell



Js x = Ps x ([Sx]o - [Sx]i )

1-7

(1.1)

The net flux of Sx can be decomposed into a unidirectional influx (Js xo→i) and a unidirectional efflux (Js x i→o). The net flux of Sx, as shown in Equation 1.1, is simply the difference between the unidirectional fluxes. Thus, the unidirectional influx is proportional to the outside concentration, the unidirectional efflux is proportional to the inside concentration, and the net flux is proportional to the concentration difference; in all cases the proportionality constant is PSx . The following equation allows the calculation of the electrochemical energy difference or electrochemical potential of Sx (ΔµSx). This parameter combines values of concentration and voltage gradients across a membrane:

ΔµSx = RT ln([Sx]i/[Sx]o) + zXF(ψi–ψo)

(1.2)

where zX is the charge of Sx, T is the absolute temperature, R is the gas constant, and F is the Faraday constant; this part of the equation defines the electrical energy difference. The term RT ln([Sx]i/[Sx]o) describes the energy (joules/mole) change as Sx moves across the membrane; it is a measure of the ­chemical energy difference. The term zXF(ψi–ψo) describes the energy change as a mole of charged particles (each with a valence of zX) moves across the membrane. The difference (ψi–ψ0) is a voltage difference across the membrane (membrane potential difference, Vm). By definition, Sx is at equilibrium when the electrochemical potential difference for Sx across the membrane is zero (ΔµSx = 0). When ΔµSx is not zero, its value represents the net driving force, causing Sx to either enter or leave the cell, provided that a pathway exists for it to cross the membrane. It is important to consider a couple of special cases for the equilibrium state equation (Equation 1.2). In the first case, when either the chemical or the electrical term in this equation is zero, for instance, when Sx is uncharged (zX = 0), as in the case of glucose, then equilibrium occurs only when [Sx] is equal at both sides of the membrane. Alternatively, when Sx is charged as in the case of the Na+, and the electrical potential difference and thus Vm are zero, equilibrium likewise occurs only when [Sx] is equal on both sides of the membrane. The second case is when neither the chemical nor the electrical term in Equation 1.2 is zero; equilibrium occurs when the two terms are equal but of opposite signs. This relationship is the Nernst equation, when ΔµSx from Equation 1.2:

Vm = EX = –(RT/zXF) ln ([Sx]i/[Sx]o)

(1.3)

Hence, the Nernst equation describes the conditions when an ion is in equilibrium across a ­ embrane. Given values for [Sx]i and [Sx]o, Sx can be in equilibrium only when the voltage m ­d ifference across the membrane equals the equilibrium potential (E X), also know as the Nernst potential [24]. Transport processes can be studied by applying some of the tools employed for enzyme kinetics analyses. Transport can be regarded as a reaction, where substrate location, and not its structure, is changed. Mathematical/graphical analyses of experimental data, similar to those used in enzyme kinetics, can be performed. Plots such as velocity versus substrate concentration (Michaelis and Menten model; Equation 1.4), as shown in Figure 1.6 and Equation 1.4, and reciprocal plots (Lineweaver–Burk) are used to characterize kinetics and the type of inhibition.

v = Vmax [Sx]/(Km + [Sx])

(1.4)

The values for Km can vary considerably from one transporter to another, and for a particular transporter with different solutes. For transport studies, the definition of Km is the solute concentration that

1-8

Cellular Metabolism

Transport rate

Vmax

½ Vmax

Km

Substrate concentration (S)

Figure 1.6  Michaelis and Menten kinetics of a transport-mediated solute diffusion process.

results in half-maximal velocity for the transport reaction. An equivalent way of stating this is that Km represents the substrate concentration at which half of the transporter is occupied by solute molecules in steady state. Hence, the constant Km can be used as a relative measure of substrate binding affinity.

1.3 The Transporter Classification (TC) System The phylogenetic and functional analyses of transporter proteins from many different organisms have provided the basis for the development of a transporter classification (TC) system [25]. In this system, permeases are classified according to both function and phylogeny. The TC system is an International Union of Biochemistry and Molecular Biology (IUBMB) approved system of nomenclature for transport protein classification. The systematic classification of solute transporters is based on several criteria including mode of transport, energy-coupling source, and molecular phylogeny. A specific five-digit TC number classifies each transporter according to five criteria. The first digit is a number referring to the class, which defines the mode of transport and energy-coupling mechanism; the second digit is a letter indicating the subclass as defined by the type of transporter and the energy ­coupling mechanism; the third digit is a number indicating the superfamily or family; the fourth digit is a number indicating a phylogenetic cluster within a family or a family within a superfamily; and the last digit is a number indicating the substrate or range of substrates transported and the polarity of transport. For example, the TC number for the galactose:H + symporter form E. coli (GalP) is 2.A.1.1.1. This number indicates that GalP is a member of the 2.A.1.1. sugar porter family from the 2.A.1. major facilitator superfamily of the 2.A. porters subclass of the 2. electrochemical potentialdriven class of transporters. The functional and phylogenetic TC system taxonomy can be accessed in the transporter classification database (TCDB). This is a Web accessible (http://www.tcdb.org) relational database containing a wealth of information about transport systems. It is a curated database that continuously updates and classifies structural, functional, evolutionary, and sequence information. The TCDB compiles information from over 10,000 references and includes approximately 3,000 different proteins organized in approximately 400 transporter families. The Web interface provides several methods for accessing data and searching the database. It also includes several bioinformatic tools designed to analyze transport proteins [26]. Table 1.2 shows an outline of TC according to permease type and energy source. There are four known classes of transporters: (1) channels, (2) porters, (3) primary active transporters, and (4) group translocators. An additional class (8) includes auxiliary transport proteins and class includes

Solute Transport Processes in the Cell

1-9

Table 1.2  Classes and Subclasses of Transporters According to the TC Systema 1.  Channels and Pores 1.A  α-Type channels 1.B  β-Barrel porins 1.C  Pore-forming toxins (proteins and peptides) 1.D  Nonribosomally synthesized channels 2.  Electrochemical Potential-Driven Transporters 2.A  Porters (uniporters, symporters, antiporters) 2.B  Nonribosomally synthesized porters 2.C  Ion-gradient-Driven energizers 3.  Primary Active Transporters 3.A  P-P-bond hydrolysis-Driven transporters 3.B  Decarboxylation-Driven transporters 3.C  Methyltransfer-Driven transporters 3.D  Oxidoreduction-Driven transporters 3.E  Light absorption-Driven transporters 4.  Group Translocators 4.A  Phosphotransfer-Driven group translocators 5.  Transmembrane Electron Flow Systems 5.A  Two-electron carriers 5.B  One-electron carriers 8.  Accessory Factors Involved in Transport 8.A  Auxiliary transport proteins 9.  Incompletely Characterized Transport Systems 9.A  Recognized transporters of unknown biochemical mechanism 9.B  Putative but uncharacterized transport proteins 9.C  Functionally characterized transporters lacking identified sequences a

Categories 6–7 are reserved for yet to be discovered novel types of transporters.

uncharacterized sequences that are homologous to transporters and characterized transporters whose sequences are not known. Categories 6–7 are reserved for yet to be discovered novel types of transporters. Figure 1.7 shows the known classes and mechanisms of solute transporters found in bacteria that include: (a) channel and pore-mediated passive diffusion; (b) carrier-mediated solute-H+ symport; (c) carrier-mediated solute-H+ symport with an external solute-recognition receptor; (d) primary active uptake ABC transporter driven by ATP hydrolysis; and (e) group translocating ­permease of the phosphoenolpyruvate:sugar phosphotransferase system (PTS). Each of these classes will be presented in detail in the following sections.

1.3.1  Channels and Pores The channel or pore is one of the simplest structures for the transport of solutes (Figure 1.7a). In this case, facilitated diffusion is not coupled to the use of metabolic energy, and hence it cannot generate concentration gradients of the transported substrate across the membrane. In this class of transporters, the solute passes by a diffusion-limited process from one side of the membrane to the other via a channel or pore that is coated by amino acyl residue moieties of the constituent protein(s) that recognizes hydrophilic, hydrophobic, or amphipathic substrates. In Gram-negative bacteria some small solutes that in principle can cross the membrane relatively without restraint may need an additional transport system to sustain a high enough flux for physiological processes. Channel type proteins, called porins, that mediate the passive transfer of solutes, perform transport of molecules such as carbohydrates, amino acids, and simple ions across the outer membrane. A bacterium like E. coli can have up to 105 copies of porins OmpF, OmpC, or PhoE,

1-10

Cellular Metabolism (a)

(b) H++ +

H

(c) H++ H+ (d)

ATP Pi + ADP

(e) PEP EI PYR

HPr IIA P~

IIB

IIC IIC

Figure 1.7  Classes and mechanisms of solute transporters. (a) Channel and pore-mediated passive diffusion. (b) Carrier-mediated solute-H+ symport. (c) Carrier-mediated solute-H+ symport with an external solute-recognition receptor. (d) Primary active uptake ABC transporter driven by ATP hydrolysis. (e) Group translocating permease of the phosphoenolpyruvate:sugar phosphotransferase system.

which tend to form a barrel of antiparallel β-sheets, consisting of trimeric complexes of identical subunits of ca. 35 kDa, with an approximate diameter of 1 nm that allows the passage for molecules up to a mass of about 600 Da. 1.3.1.1  α-Type Channels This class of channel proteins is composed mainly of α-helical spanners (sometimes β-strands play a part in the channel) that catalyze the movement of solutes using transmembrane aqueous pores by an energy-independent mechanism and are found in bacteria and eukaryotes. 1.3.1.2  β-Barrel Porins These porin proteins are mainly β-barrels constituted by β-strands in the transmembrane region, which also allow the transport of solutes by an energy-independent mechanism. They are found in the outer membranes of Gram-negative bacteria, mitochondria, and plastids.

Solute Transport Processes in the Cell

1-11

1.3.1.3  Pore Forming Toxins (Proteins and Peptides) This category of transmembrane pores includes proteins or peptides ribosomally synthesized by one cell and secreted for insertion into the membrane of another cell. They are composed mainly of β-strands. The pores formed by these proteins allow the free flow of electrolytes and other small molecules across the membrane of the host, or they may allow entry into the target cell cytoplasm of a toxin protein that ultimate kills or controls the cell. 1.3.1.4 Nonribosomally Synthesized Channels These oligomeric transmembrane ion channels are chains of L- and D-amino acids, or small polymers of hydroxylactate or β-hydroxybutirate, in which the arrangement of the pore structure is stimulated by voltage changes, and are made by bacteria and fungi as agents of biological warfare that confer a selective advantage.

1.3.2 Electrochemical Potential-Driven Transporters Chemical, light, or electrochemical energy is used by the cells to transport and accumulate solutes inside the cell. Several transport phenomena are driven by electrochemical potential gradients, such as proton and sodium gradients. Concentration gradients of solutes power transporter-mediated facilitated diffusion; it does not require the expenditure of metabolic energy in the form of ATP. This mechanism permits the transfer of solutes across a membrane against a concentration gradient. This type of transport is also known as secondary transport and is catalyzed by uniporters, symporters, and antiporters (Figure 1.7b and c). In contrast to transporters driven by ATP, electrochemical transporters are relatively simple in composition, generally consisting of a single protein that transverses the membrane with several loops. 1.3.2.1  Porters (Uniporters, Symporters, and Antiporters) Some sugars, amino acids, nucleosides, and small molecules, such as Na+, are transported by uniporter proteins, which move one solute across a membrane down a concentration gradient from an area of greater concentration to one of lesser concentration. Selective conformational changes are induced by interactions between the solute and the uniporter, which enables the uniporter to transport the solute across the cytoplasmic membrane. Some sugars, amino acids, and ions (e.g., sulfate and phosphate) are co-transported by symporter proteins, which use the proton motive force to move a solute against a concentration gradient, i.e., symporters simultaneously transport two solutes across the membrane in the same direction. Antiport involves a tightly coupled process where two or more solutes are transported in opposite directions. Na + can be transported by antiporter proteins. In this instance, a gradient of protons generates the potential energy, and simultaneously the solute (Na+) is transported trough the membrane in the opposite direction against a concentration gradient. 1.3.2.2 Nonribosomally Synthesized Porters These transmembrane porters are peptides or small polymers of nonpeptide nature. They complex ­solutes (like a cation) in their hydrophilic interior and make possible the translocation of the complex across the membrane by exposing its hydrophobic exterior and moving from one side of the membrane bilayer to the other. Transport is electrophoretic if the porter in the uncomplexed form can cross the membrane and electroneutral (one charged solute is exchanged for another) if only the complexed form can cross the membrane. 1.3.2.3  Ion-Gradient-Driven Energizers This is a family of auxiliary proteins, like the TonB family, that mediate active transport using an outer membrane receptor, which is energized to accumulate its solutes inside the periplasm against large

1-12

Cellular Metabolism

IICBGlc

IICGal IIDGal

IICBMal

IIB'BCFru

IIBGal IIAGal ~P

IIAGlc ~P

PEP

EI

Pyruvate

EI

IICMan IIDMan IIABMan

IICBAMtl

IIAMHFru ~P

Hpr ~P ~P

Hpr

Figure 1.8  The phosphoenolpyruvate:sugar phosphotransferase system from Escherichia coli. General ­energy-coupling proteins and some sugar transporting complexes are shown. Glc, glucose; Mal, maltose; Gal, galactosamine; Fru, fructose; Man, mannose; and Mtl, mannitol.

concentration gradients. These energizers make use of protons or sodium ions fluxes, i.e., the proton motive force, through themselves to energize outer membrane receptors or porins. Conformational changes of receptors allow electrophoretic transport of protons.

1.3.3  Primary Active Transporters These systems consist of transporters that use a primary source of energy to drive the active transport of a solute against a concentration gradient. Primary energy sources known to be coupled to transport include chemical, electrical, and light. 1.3.3.1  P-P-Bond-Hydrolysis-Driven Transporters These transport systems hydrolyze the diphosphate bond of inorganic pyrophosphate, ADP, ATP, or another nucleoside triphosphate, to drive the active uptake and/or extrusion of a solute or solutes. The transport protein may or may not be transiently phosphorylated, but the substrate is not phosphorylated during the process. This subclass is the most abundant of the primary active transporters and comprises three superfamilies and 18 families distributed among all prokaryotic and eukaryotic organisms (Table 1.3). 1.3.3.2  Decarboxylation-Driven Transporters This subclass includes transport systems that drive ion uptake or extrusion by decarboxylation of a cytoplasmic substrate. The only family included in this subclass is the Na+ -transporting carboxylic acid decarboxylase (NaT-DC) system that catalyzes decarboxylation of carboxilic substrates such as oxaloacetate, methylmalonyl-CoA, glutaconyl-CoA, and malonate, respectively, and use the energy released to drive extrusion of one or two sodium ions (Na+) from the cell cytoplasm. The α-subunits of these transporters are biotin containing multisubunit enzymes and are currently thought to be restricted to bacteria. However, on the basis of the homology of the α-δ-subunits to proteins coded by Archaea genomes, it is proposed that NaT-DC family members may also be present in this kingdom. 1.3.3.3  Methyltransfer-Driven Transporters The Na + transporting methyltetrahydromethanopterin:coenzyme M methyltransferase (NaTMMM) of the Archaea Methanobacterium thermoautotrophicum is the only characterized

1-13

Solute Transport Processes in the Cell Table 1.3  Overview of the Superfamilies and Families of the P-P-Bond-Hydrolysis-Driven Transporters Superfamily or Familya

Mechanism: Solute(s) Tranported

3.A.1 ATP-binding cassette (ABC) superfamily

Bacterial and archaeal ABC-type uptake permeases: carbohydrates, polar-, hydrophobicaminoacids, peptide/opine/nickel, sulfate, phosphate, molybdate, phosphonate, ferric iron, poliamine/opine/phosphonate, quaternary amine, vitamin B12, iron chelate, manganese/ zinc/iron chelate, nitrate/nitrite/chelate, taurine, cobalt, thiamin, iron, Fe3+, nickel. Bacterial ABC-type efflux (exporter) permeases: capsular polysaccharide, lipooligosaccharide, lipopolysaccharide, teichoic acid, drug, lipid, heme, β-glucan, protein, peptide, glycolipid, Na+, microcin B17, drug/siderophore, drug resistance ATPase. Eukaryotic ABC-type efflux (exporter) permeases: multidrug and pleitropic drug resistance, cystic fibrosis transmembrane conductance, peroxysomal fatty Acyl CoA, eye pigment precursor, a-factor sex pheromone, conjugate transporter, MHC peptide, heavy metal, cholesterol/phospholipid/retinal, mitochondrial peptide.

3.A.2 and 3.A.3 F- and P-typeATPase superfamilies 3.A.4 to 3.A.18 Families

H+- or Na+-translocation

a

Arsenite-antimonite efflux; diverse secretory pathways; mitochondrial-, chloroplast envelop protein-, H+ - and septal DNA-translocase systems; bacterial competence-related DNA transformation transporter; filamentous phage-, fimbrilin- and the nuclear mRNA exporters; the endoplasmic reticular retrotranslocon; the phage T7 injectisome.

multisubunit protein family of this subclass. Analysis of the coding genes revealed eight contiguous genes organized in an operon. The majority of the encoded subunits are probably integral membrane proteins. 1.3.3.4 Oxidoreduction-Driven Transporters This subclass includes those transport systems that drive the transport of a solute (proton or ion) energized by the exothermic flow of electrons from a reduced substrate to an oxidized substrate. Transporters included in this subclass comprise two superfamilies and seven families (Table 1.4). These transporters, are found in bacteria, Archaea, eukaryotic mitochondria, and chloroplast; however, some families are group restricted. 1.3.3.5  Light Absorption-Driven Transporters The members of these families catalyze light-driven ion translocation across microbial cytoplasmic membranes or serve as light receptors. There are two families included: the 3.E.1 ion-translocating microbial rhodopsin (MR) and the 3.E.2 photosynthetic reaction center (PRC) families.

1.3.4  Group Translocators 1.3.4.1  Phosphotransfer-Driven Group Translocators This class of transporters includes the phosphoenolpyruvate:sugar PTS (Figure 1.7e). This protein system is widespread in bacteria but absent in Archaea, and eukaryotic organisms [27]. The PTS transport mechanism involves the transport and phosphorylation of several sugars. This is a unique characteristic of the group translocator class of transporters. The product of the reaction is a sugar-phosphate that will subsequently enter a catabolic pathway. The system is composed of the soluble and nonsugar-specific protein components Enzyme I (EI) and the heat-stable or phosphohistidine carrier protein (HPr). The first step of the PTS mechanism involves autophosphorylation by PEP of EI at a histidine residue. In the second step, EI transfers the phosphoryl group to a histidine residue in the HPr. Then, the HPr transfers the phosphoryl group to proteins called enzymes IIA and IIB, which are part of each sugar-specific PTS complex. The final component of this system, IIC (in some cases also IID), is an integral membrane protein permease that recognizes and

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Cellular Metabolism

Table 1.4  General Characteristics of the Oxidoreduction-Driven Transporter Families Family/Superfamily 3.D.1. Proton-translocating NADH dehydrogenase (NDH) family 3.D.2 Proton-translocating transhydrogenase (PTH) family 3.D.3 Proton-translocating quinol:cytochrome c reductase (QCR) superfamily 3.D.4 Proton-translocating cytochrome oxidase (COX) superfamily

Generalized Transport Reaction NADH + ubiquinone + 4H+ (in) → NAD+ + ubiquinol + 4H+ (out), or NADH + ubiquinone + 2Na+ (in) → NAD+  + ubiquinol + 2H+ (out) → NADPH + NAD+ + H+ (in) NADP+ + NADH + H+ (out) ←  quinol (QH2) + 2 cytochrome c (ox) + 2H+ (in)→quinone (Q) + 2 cytochrome c (red) + 4H+ (out) Reaction catalyzed by cytochrome c (Cyt c) oxidases: 2Cyt c (red) + 1/2 O2 + 6H + (in)→2Cyt c (ox) + H2O + 4H+ (out) Reaction catalyzed by quinol oxidases is: quinol + 1/2 O2 + 4H+ (in)→quinone + H2O + 4H+ (out)

3.D.5 Na+ -translocating NADH:quinone dehydrogenase (Na-NDH) family

NADH + quinone + nNa+ (in)→NAD+ + quinol + nNa+ (out)

3.D.6. Putative ion (H + or Na+)translocating NADH:ferredoxin oxidoreductase (NFO) family 3.D.7 H2:Heterodisulfide oxidoreductase (HHO) family

NADH + oxidized ferredoxin + n(H+ or Na+) (out)→NAD+ + reduced ferredoxin + n(H+ or Na+) (in).

3.D.8 Na+ or H+ pumping formyl methanofuran dehydrogenase (FMF-DH) family 3.D.9 The H+ -translocating F420H2 dehydrogenase |(F420H2DH) family

Reactions which are coupled to H+ translocation (extrusion): (1) H2 + 2-hydroxyphenazine + H+ (in)→dihydro-2hydroxyphenazine + H+ (out) (2) dihydro-2-hydroxyphenazine + CoM-S-S-CoB + H + (in)→2hydroxyphenazine + HS-CoM + HS-CoB + H+ (out) → CO  + MF + H  + 2Na+ or 2H+ (out) Formyl-MF + 2Na+ or 2H+ (in) ←  2

2

→ oxidized acceptor (2e–) + 1H+ (out). F420H2 reduced donor (2e–) + 1H+ (in) ←  cofactor

transports the sugar molecule, which is phosphorylated by component IIB (Figure 1.8). The PTS of each organism usually includes several different enzyme II complexes, each one usually specific for single sugar substrates. In the case of E. coli, there are 26 different identified enzyme II complexes. It should be noted that there is significant diversity with regard to domain composition for the sugar complexes. As it can be seen in Figure 1.8, the maltose/glucose PTS complex form of E. coli lacks the IIA domain. Presumably, the IIA protein from the glucose complex (IIAGlc) is involved in the phosphorylation of the IIBMal domain. Also evident in Figure 1.8 is the frequent occurrence of protein fusions among enzyme II domains. Fusions including IIA-IIB, IIB-IIC, and IIA-IIBIIC, among others, have been observed in different organisms. Analyses of bacterial genomes have revealed a high diversity with regard to PTS composition. Some bacteria have a single enzyme II complex, whereas others have several dozens. These studies have also revealed that some bacterial species completely lack genes encoding PTS components, whereas others have EI and/or HPr proteins but none of the enzyme II complexes. It is possible that in these cases the PTS proteins have a regulatory role [28]. PEP-dependent phosphorylation by PTS results in a tight linkage between sugar transport and its subsequent metabolism (Figure 1.9). PEP can be considered a link between the Embden–Meyerhof– Parnas (EMP) glycolytic pathway and the PTS, together forming a phosphorylation circuit. PEP is a highly connected compound in the central metabolic network of all organisms. This metabolite is a precursor in several biosynthetic pathways and also participates directly in energy-generating reactions such as substrate-level phosphorylation of ADP or indirectly as a precursor of acetyl-CoA. The relative carbon flux originating from the PEP node into the different metabolic pathways has been determined experimentally. When E. coli grows in minimal medium containing glucose as the

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Solute Transport Processes in the Cell Glucose IICBGlc

IIAGlc ~P

G6P F6P

HPr ~P F1,6BP

EI ~P

GAP + DHA Cell wall

PEP + PEP

Aromatic compounds

Acetate PYR

AcCoA CIT ICIT OAA AKG

MAL FUM

SUCCoA SUC

Figure 1.9  Central pathways related to glucose transport and metabolism in Escherichia coli. Dashed lines indicate more than one biochemical reaction.

carbon source, the PTS consumes 50% of the available PEP, whereas the reactions leading to the synthesis of oxaloacetate, pyruvate, cell wall components, and aromatic compounds consume approximately 16%, 15%, 16%, and 3%, respectively [29,30]. From these data, it is evident that PTS is a cellular activity having a major influence on the PEP/PYR ratio and the carbon flux distribution originating from these two metabolic nodes. It can be expected, then, that modification or elimination of PTS components should have a significant impact on central metabolism. PTS modification or its functional replacement has been a strategy applied in E. coli to improve the properties of production strains. Expression of PEP-independent uptake and phosphorylation activities in strains devoid of PTS activity has resulted in significant improvements in productivity and product yield for several classes of metabolites [31]. A common feature of most free-living bacteria is their capacity to select, from a mixture of carbon sources, the one that affords the highest growth rate. This response is known as carbon catabolite

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Cellular Metabolism

repression (CCR) and it is identified by the inhibition of sugar transport capacity, enzyme activities, and gene expression by the presence of a rapidly metabolizable carbon source [32]. Glucose repression is another name usually employed to describe this phenomenon, although other PTS sugars can also exert CCR. This event is responsible for diauxic growth, observed when bacteria are cultured in media containing more than one carbon source. In the case of E. coli, the IIAGlc protein has a central role in CCR. When glucose is present in the medium, EI, Hpr, and IIAGlc are present mainly in a nonphosphorylated state, since the phosphoryl group from PEP is transferred, via IIBCGlc, to glucose. In this condition, IIAGlc binds to various nonPTS permeases, inhibiting the uptake of non-PTS sugars (Figure 1.10a). Dephosphorylated IIAGlc can also bind to the enzyme glycerol kinase (GK), inhibiting its activity [33]. Proteins EI and Hpr also have regulatory roles that are controlled by phosphorylation. Dephosphorylated EI has been shown to bind to the chemotaxis protein CheA, inhibiting its autophosphorylating activity and thus causing smooth swimming behavior [34]. When it is dephosphorylated, Hpr activates the enzyme glycogen phosphorylase (GP) [35]. The protein IIBCGlc plays a role as regulator of PTS and thus, indirectly, in CCR. This protein interacts with the transcriptional repressor Mlc that regulates genes ptsHI, ptsG, mlc, manXYZ, and malT. Glucose presence causes dephosphorylation of IIBGlc under this condition; it binds Mlc, thus relieving its repression from genes under its control. The net effect of this response is an increased expression of the general PTS enzymes and those involved in glucose, mannose, and maltose transport [36]. If glucose is absent from the medium, IIAGlc and IIBGlc will be mainly in their phosphorylated states (Figure 1.10b). In this condition, IIAGlc~P binds to the enzyme adenylate cyclase (AC), activating its cAMP biosynthetic capacity. Therefore, cAMP concentration increases in the cell, binding to the cAMP receptor protein (CRP) and causing the induction of catabolite-repressed genes [37]. It should be noted that AC displays a basal level of activity even in the absence of IIAGlc~P activation. Therefore, a low level of cAMP is present in the cell when growing on glucose. In the absence of glucose, IIBGlc~P loses its capacity to bind Mlc. Thus, this regulatory protein is free to bind to its target operator sequences, causing repression of genes involved in glucose uptake. When Hpr is phosphorylated, it binds to and activates BglG, a transcriptional activator of the bgl operon that encodes proteins involved in β-glucosidic sugars uptake and utilization [38]. PTS forms part of a complex regulatory network involved in coordinating cellular processes related to the cell’s capacity to select, transport, and metabolize a large number of carbon sources [39]. Therefore, it can be expected that direct modification of PTS components will have wide-ranging effects on the cell’s physiology. As a result of CCR, E. coli displays sequential sugar consumption when it is grown in media containing sugar mixtures, like those generated from lignocellulose hydrolyzates. The simultaneous consumption of sugars present in a mixture would be advantageous in a fermentative production process, since this would eliminate diauxic growth, resulting in reduced operating time and increased productivity. Modification of components from the glucose PTS complex in E. coli has resulted in disruption of CCR. This strategy has been applied to improve strains for the production of ethanol and L-lactic using mixtures of glucose, arabinose, and xylose as carbon sources [11,40,41]. It is expected that similar strategies should also improve the characteristics of production strains derived from other ­bacterial species that have PTS.

1.3.5 Transmembrane Electron Flow Systems These systems are involved in the catalytic flow of electrons across a biological membrane, from donors localized to one side of the membrane to acceptors localized on the other side. These systems contribute to or subtract from the membrane potential, depending on the direction of electron flow, and are important elements in cellular energetics. According to the TCDB, members of this class are grouped in two subclasses: (1) transmembrane two-electron transfer carriers and (2) transmembrane one-electron transfer carriers (see Table 1.5).

EI

PEP

–

EI

Pyruvate

Hpr ~P

Hpr

+

–

GK

GP

~P

IIAGlc

IIAGlc

AC

–

Glucose

–35 –10 Mlc-repressed genes

G6P

Mlc IICB

Glc

–10 –35

EI

PEP +

~P

PTS-sugars~P

BglG

EI

Pyruvate

Catabolite-repressed genes

+

CRP

S -PT Non eases m r pe

Hpr ~P

Hpr

cAMP

ATP

PTS-sugars

PTS ases r e p me

IIAGlc

~P IIAGlc

+

AC

Glc Mlc IICB

Mlc-repressed genes

Mlc

P

Figure 1.10  Carbon catabolic repression mechanisms in Escherichia coli. Regulatory interactions of PTS components in presence (a) or absence (b) of glucose.

~P

–10 –35 Catabolite-repressed genes

CRP

(b)

~

CheA

(a)

Solute Transport Processes in the Cell 1-17

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Cellular Metabolism

Table 1.5  Overview of the Subclasses of the Transmembrane Electron Flow Systems Class/Subclass/Family Transmembrane two-electron transfer carriers. 5.A.1. Disulfide bond oxidoreductase D (DsbD)

Mechanism: Solute(s) Transported Catalyses an essentially irreversible reaction due to the fact that electrons flow down their electrochemical gradient from inside the bacterial cell (negative inside) to outside the cell (positive outside). In order to reverse the reaction, electrons are transferred from dithiol proteins in the periplasm to an electron acceptor in the cytoplasm. The overall vectorial electron transfer reaction catalyzed by DsbD is 2 e–cytoplasm→2 e–periplasm

5.A.2. Disulfide bond oxidoreductase B (DsbB)

DsbB family transfers electrons across the bacterial cytoplasmic membrane from periplasmic dithiol proteins to quinones in the membrane. The reduced quinones then reduce cytoplasmic electron acceptors such as O2 via cytochrome oxidases (cytochrome d and cytochrome o, respectively, in E. coli) as well as nitrate via nitrate reductase and fumarate via fumarate reductase. The generalized vectorial electron transfer reaction catalyzed by DsbB is 2 e–cytoplasm→2 e–periplasm

5.A.3. Prokaryotic molybdopterincontaining oxidoreductase (PMO)

The membrane-bound nitrate reductase-A (NR-A) (contains Mo-molybdopterin guanine dinucleotide) and formate dehydrogenase (FDH) employ a redox loop to couple quinol oxidation to the equivalent of proton translocation, thereby generating a proton motive force (pmf) during anaerobic respiration. The net transmembrane electron transfer reactions for NR-A and FDH, and probably other homologous enzymes, are a. 2e– (out)→2e– (in) (NR-A) b. 2e– (in)→2e– (out) (FDH)

Transmembrane one-electron transfer carriers: 5.B.1 gp91phox phagocyte NADPH oxidaseassociated cytochrome b558 (Phox)

The human phagocyte cytochrome b558 is a heterodimeric complex consisting of a heavy (β) chain (gp91phox) and a light (α) chain (p22phox) as well as several auxiliary subunits. gp91phox is the terminal component of a respiratory chain that transfers single electrons from cytoplasmic NADPH to O2 on the external side of the plasma membrane generating superoxide. Its activity is electrogenic, causing depolarization of the membrane potential on the negative inside. The overall electron transfer reactions catalyzed by gp91phox and some of its homologues are: 1. electron (in)→electron (out) 2. O2 (out) + electron (in)→O2– (superoxide) (out)

5.B.2 Eukaryotic cytochrome b561 (Cytb561)

Bovine cytochrome b561 is a six TMS protein present in secretory vesicles (i.e., adrenal chromaffin granules) that contain catecholamines and amidated peptides. It supplies electrons from cytoplasmic L-ascorbate to intravesicular enzymes such as dopamine β-hydroxylase and α-peptide amidase. Transmembrane electron flow reactions catalyzed by Cytb561 family members are 1. L-ascorbate (cytosol) + enzyme (oxidized) (synaptic vesicle)→dehydroascorbate (cytosol) + enzyme (reduced) (synaptic vesicle) 2. L-ascorbate (cytosol) + 2Fe3+ (intestinal lumen)→dehydroascorbate (cytosol) + 2Fe2+ (intestinal lumen).

5.B.3 Geobacter nanowire electron transfer (G-NET)

Geobacter and other bacteria from diverse kingdoms have the capacity to transfer electrons from cytoplasmic electron donors to extracellular substances. Geobacter utilizes “nanowires” distantly related to type IV pili (fimbriae) to transfer electrons to iron oxide (Fe2O3; rust) to generate soluble Fe2 + and solid magnetite. An electron pair is transferred from NADH to menaquinone, and then single electrons are transferred through the chain to iron oxide via an inner membrane (IM), periplasm, outer membrane (OM) pathway as follows: NADH → NADHDH (3.D.1.4.1) → menaquinone → MacA (IM) → PpcA (periplasm) →  OmcB (a trans OM protein) → OmcE (outer surface of the OM) → OmcS (outer surface of the OM) → pilin → Fe2O3.

5.B.4 Plant photosystem I supercomplex (PSI)

Water, the electron donor for photosynthesis, is oxidized to O2 and four protons by PSII. The electrons that have been extracted from water are shuttled through a quinone pool and the cytochrome b6f complex to plastocyanin—a small, soluble, copper-containing protein. Solar energy that has been absorbed by PSI induces the translocation of an electron from plastocyanin at the inner face of the membrane (thylakoid lumen) to ferredoxin on the opposite side (stroma).

Solute Transport Processes in the Cell

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References 1. Paulsen, I. T., Sliwinski, M. K., and Saier, M. H. Jr. Microbial genome analyses: global comparisons of transport capabilities based on phylogenies, bioenergetics and substrate specificities. J. Mol. Biol., 277, 573, 1998. 2. Saier, M. H. Jr. Computer-aided analyses of transport protein sequences: gleaning evidence concerning function, structure, biogenesis, and evolution. Microbiol. Rev. 58, 71, 1994. 3. Díaz, E. et al. Biodegradation of aromatic compounds by Escherichia coli. Microbiol. Mol. Biol. Rev., 65, 523, 2001. 4. Bruckner, R. and Titgemeyer, F. Carbon catabolite repression in bacteria: choice of the carbon source and autoregulatory limitation of sugar utilization. FEMS Microbiol. Lett., 209, 141, 2002. 5. Ferenci, T. Adaptation to life at micromolar nutrient levels: the regulation of Escherichia coli glucose transport by endoinduction and cAMP. FEMS Microbiol. Rev., 18, 301, 1996. 6. Hahn-Hägerdal, B. et al. Metabolic engineering of Saccharomyces cerevisiae for xylose utilization. Adv. Biochem. Eng. Biotechnol., 73, 53, 2001. 7. Flores, N. et al. Pathway engineering for the production of aromatic compounds in Escherichia coli. Nat. Biotechnol., 14, 620, 1996. 8. Yi, J. et al. Altered glucose transport and shikimate pathway product yields in E. coli. Biotechnol. Prog., 19,1450, 2003. 9. Lin, H., Bennett, G. N., and San, K. Y. Metabolic engineering of aerobic succinate production ­systems in Escherichia coli to improve process productivity and achieve the maximum theoretical succinate yield. Metab. Eng., 7, 116, 2005. 10. Hernández-Montalvo, V. et al. Expression of galP and glk in a Escherichia coli PTS mutant restores glucose transport and increases glycolytic flux to fermentation products. Biotechnol. Bioeng., 83, 687, 2003. 11. Dien, B. S., Nichols, N. N., and Bothast, R. J. Fermentation of sugar mixtures using Escherichia coli catabolite repression mutants engineered for production of L-lactic acid. J. Ind. Microbiol. Biotechnol., 29, 221, 2002. 12. de Anda, R. et al. Replacement of the glucose phosphotransferase transport system by galactose permease reduces acetate accumulation and improves process performance of Escherichia coli for recombinant protein production without impairment of growth rate. Metab. Eng., 8, 281, 2006. 13. Palsdottir, H. and Hunte, C. Lipids in membrane protein structures. Biochim, Biophys, Acta, 1666, 2, 2004. 14. Cabeen, M. T. and Jacobs-Wagner, C. Bacterial cell shape. Nat. Rev. Microbiol., 3, 601, 2005. 15. Desvaux, M. et al. Protein cell surface display in Gram-positive bacteria: from single protein to macromolecular protein structure. FEMS Microbiol. Lett., 256, 1, 2006. 16. Volkman, J. K. Sterols in microorganisms. Appl. Microbiol. Biotechnol., 60, 495, 2003. 17. Singer, S. J. and Nicholson, G. L. The fluid mosaic model of the structure of cell membranes. Science, 175, 720, 1972. 18. Matsumoto, K. et al. Lipid domains in bacterial membranes. Mol. Microbiol., 6, 1110, 2006. 19. Heathcock, C. H. et al. Stereostructure of the archaebacterial C40 diol. Science, 229, 862, 1985. 20. Tornabene, T. G. and Langworthy, T. A. Diphytanyl and dibiphytanyl glycerol ether lipids of methanogenic archaebacteria. Science, 203, 51, 1979. 21. Moat, A. G., Foster, J. W., and Spector, M. P. Microbial Physiology. Wiley-Lyss. New York, 2002, ­chapter 7, Cell structure and function. 22. Fu, D. et al. Structure/function relationships in OxlT, the oxalate-formate transporter of Oxalobacter formigenes. J. Biol. Chem., 276, 8753, 2001. 23. Ames, J. F. Bacterial periplasmic transport systems: structure, mechanism, and evolution. Annu. Rev. Biochem., 55, 397, 1986. 24. Suzuki, H. et al. Probing the transmembrane potential of bacterial cells by voltage-sensitive dyes. Anal. Sci., 19, 1239, 2003.

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Cellular Metabolism

25. Saier, M. H. Jr. A functional-phylogenetic classification system for transmembrane solute transporters. Microbiol. Mol. Biol. Rev., 64, 354, 2000. 26. Saier, M. H. Jr., Tran, C. V., and Barabote, R. D. TCDB: the Transporter Classification Database for membrane transport protein analyses and information. Nucleic Acids Res., 34, D181, 2006. 27. Postma, P. W., Lengeler, J. W., and Jacobson, G. R. Phosphoenolpyruvate: carbohydrate phosphotransferase systems. In Escherichia coli and Salmonella. Cellular and Molecular Biology, Neidhardt, F. C., Ed. American Society for Microbiology, Washington, DC, 1996. 28. Barabote R. D. and Saier M. H. Jr. Comparative genomic analyses of the bacterial phosphotransferase system. Microbiol. Mol. Biol. Rev., 69, 608, 2005. 29. Valle, F. et al. Basic and applied aspects of metabolic diversity: the phosphoenolpyruvate node. J. Ind. Microbiol., 17, 458, 1996. 30. Flores, S. et al. Analysis of carbon metabolism in Escherichia coli strains with an inactive phosphotransferase system by 13C labeling and NMR spectroscopy. Metab. Eng., 4, 124, 2002. 31. Gosset, G. Improvement of Escherichia coli production strains by modification of the phosphoenolpyruvate:sugar phosphotransferase system. Microb. Cell. Fact., 4, 14, 2005. 32. Saier, M. H., Ramseier, T. M., and Reizer, J. Regulation of carbon utilization. In Escherichia coli and Salmonella. Cellular and Molecular Biology, Neidhardt, F. C., Ed. American Society for Microbiology, Washington, DC, 1996. 33. Novotny, M. J. et al. Allosteric regulation of glycerol kinase by enzyme IIIglc of the phosphotransferase system in Escherichia coli and Salmonella typhimurium. J. Bacteriol., 162, 816, 1985. 34. Lux, R. et al. Coupling the phosphotransferase system and the methyl-accepting chemotaxis protein-dependent chemotaxis signaling pathways of Escherichia coli. Proc. Natl. Acad. Sci., 92, 11583, 1995. 35. Seok, Y. J. et al. Regulation of E. coli glycogen phosphorylase activity by HPr. J. Mol. Microbiol. Biotechnol., 3, 385, 2001. 36. Plumbridge, J. Regulation of gene expression in the PTS in Escherichia coli: the role and interactions of Mlc. Curr. Opin. Microbiol., 5, 187, 2002. 37. Korner, H., Sofia, H. J., and Zumft, W. G. Phylogeny of the bacterial superfamily of Crp-Fnr transcription regulators: exploiting the metabolic spectrum by controlling alternative gene programs. FEMS Microbiol. Rev., 27, 559, 2003. 38. Görke, B. and Rak, B. Catabolite control of Escherichia coli regulatory protein BglG activity by antagonistically acting phosphorylations. EMBO J., 18, 3370, 1999. 39. Saier, M. H. Jr. and Reizer, J. The bacterial phosphotransferase system: new frontiers 30 years later. Mol. Microbiol., 13, 755, 1994. 40. Lindsay, S. E., Bothast, R. J., and Ingram, L. O. Improved strains of recombinant Escherichia coli for ethanol production from sugar mixtures. Appl. Microbiol. Biotechnol., 43, 70, 1995. 41. Hernández-Montalvo, V. et al. Characterization of sugar mixtures by an Escherichia coli mutant devoid of the phosphotransferase system. Appl. Microbiol. Biotechnol., 57, 186, 2001.

2 Catabolism and Metabolic Fueling Processes 2.1 2.2 2.3 2.4

Introduction ���������������������������������������������������������������������������������������2-1 Classification of Organisms �������������������������������������������������������������2-2 Thermodynamics of Fueling Processes..........................................2-2 Products of Fueling Processes �������������������������������������������������������� 2-4

2.5

Redox Potentials and Mobile Electron Carriers........................... 2-9

2.6

Examples of Catabolic Processes in Different Organisms.........2-14

Precursor Metabolites  •  ATP  •  NADPH

Redox Potentials  •  Mobile Electron Carriers  •  Quinones •  Flavoproteins  •  Iron-Sulfur Proteins  •  Cytochromes

Olubolaji Akinterinwa and Patrick C. Cirino Pennsylvania State University

Photosynthesis  •  The Central Pathways  •  Anaplerotic and Peripheral Reactions  •  Fermentation  •  Oxidative Phosphorylation •  Key Parameters Influencing Regulation of Catabolic Pathways

2.7 Concluding Remarks ��������������������������������������������������������������������� 2-30 Acknowledgment ��������������������������������������������������������������������������������������� 2-30 References ��������������������������������������������������������������������������������������������������� 2-30

2.1  Introduction Comparative DNA and ribosomal RNA sequencing provides evidence of three phylogenetically distinct domains of organisms: bacteria, archaea, and eukarya [1]. Although all domains are believed to have originated from a common ancestor [1,2], there is an amazing diversity of mechanisms by which these different life forms harness energy from their environment to fuel essential metabolic processes that enable survival. This metabolic diversity is a testament to the power of evolution and the adaptability of life. Metabolism can be defined as the sum of processes involved in energy conversions in the cell. They regulate cellular conditions such that a state of metabolic homeostasis, i.e., a stable supply of energy and metabolites, is maintained. Metabolic processes are organized into complex sequences of controlled chemical reactions referred to as metabolic pathways, and many different pathways are responsible for nutrient processing, energy acquisition, and energy conversion in the cell. These metabolic processes can be broadly categorized as catabolic and anabolic. Catabolic processes are responsible for the production of energy, reducing power, and precursor molecules. They usually involve the breakdown of complex molecules into simpler molecules. In contrast, anabolic processes consume energy, reducing power and precursor molecules for the synthesis of complex biomolecules such as proteins, nucleic acids, and membranes. This chapter provides an overview of the predominant catabolic/fueling metabolic processes found in nature and particularly in microorganisms. Fueling reactions collectively represent the most diverse of all processes within the cell [3,4]. Since catabolic mechanisms vary widely from one organism to another, we emphasize major pathways and products, energy yields and efficiency of energy conversion. The wellstudied metabolism of Escherichia coli serves to demonstrate many of the catabolic paradigms described. 2-1

2-2

Cellular Metabolism

2.2  Classification of Organisms Catabolic processes require an initial source of energy in the form of light or chemical energy. Organisms that utilize light energy directly are called phototrophs while those that utilize chemical energy (via chemical reactions) are called chemotrophs. An overall mutualistic relationship exists between phototrophs and chemotrophs such that chemotrophs primarily derive chemical energy from substrates ultimately originating from phototrophic metabolism via photosynthesis. During photosynthesis, light energy (usually solar energy) is converted to chemical energy while CO2 is captured. In addition to requiring energy, catabolic processes also require reducing power (electrons) and carbon. The sources of electrons and carbon are used as parameters to further classify organisms as lithotrophs or organotrophs and autotrophs or heterotrophs, respectively. Lithotrophs obtain ­reducing equivalents from inorganic sources while organotrophs obtain reducing equivalents from organic sources. Autotrophs utilize CO2 as a carbon source or initial substrate while heterotrophs utilize ­carbon compounds other than CO2. Sugars such as glucose, xylose, and maltose are examples of organic sources of electrons and carbon. Examples of inorganic electron sources include H2, H2O, H2S, NH3, S, S2O32− , and Fe. A variety of combinations of energy, electron, and carbon sources is used by organisms, as shown in Table 2.1. Organisms can also be classified based on their ability to utilize oxygen for respiration or their sensitivity to oxygen. Strict/obligate aerobes require oxygen for respiration and cannot survive in the absence of oxygen. Strict/obligate anaerobes such as Clostridium spp. are sensitive to oxygen or reactive oxygen species, cannot grow in the presence of oxygen, and instead grow fermentatively or respire anaerobically. Facultative aerobes and anaerobes are more flexible with their oxygen requirements and can survive in the absence or presence of oxygen. Examples of facultative aerobes include E. coli and Bacillus anthracis. Aerotolerant anaerobes do not respire on oxygen but are able to grow in its presence.

2.3 Thermodynamics of Fueling Processes Fueling reactions essentially convert energy extracted from a substrate into a form that is more readily available and useful for driving anabolic processes. The energy yield of a fueling process depends on the substrate being decomposed and is quantified by the change in Gibbs free energy (G) for the conversions involved in the process: ∆G = Gproducts–Greactants

(2.1)

Fueling reactions which are exergonic (energy releasing) have negative ∆G values while those which are endergonic (energy consuming) have positive ∆G values. An estimate of ∆G for a chemical reaction can be obtained from the standard Gibbs free energy (G'0) of substrates which have been reported in literature. The standard Gibbs free energy change (∆G'0) applies to reactions occurring at standard temperature (T = 298K), and pH = 7, with starting substrate concentration(s) of 1 M. The relationship between the ∆G'0 and the actual ∆G of a reaction is:

ΔG=ΔG ′ 0 +RT lnQ

(2.2)

where Q is the mass action ratio, defined as the ratio of activities of products to reactants, and in the case of dilute aqueous solutions is often estimated as the ratio of concentrations of products to reactants [5]. R is the gas constant (R = 8.314 J/mole K) and T is the absolute temperature (K). For reactions at equilibrium, ∆G = 0, so that:

ΔG ′ 0 = - RT lnK eq

(2.3)

where Keq is the equilibrium constant. Fueling processes are characterized by high-energy group transfer reactions which promote the energetic favorability of substrate decomposition. These high-energy group transfers are typically in

2-3

Catabolism and Metabolic Fueling Processes Table 2.1  Common Classifications of Organisms Basis of Classification

Designation

Description

Source of carbon i. Autotroph ii. Heterotroph

Utilizes CO2 as source of carbon Utilizes organic molecules such as sugars as source of carbon

i. Organotroph ii. Lithotroph

Obtains electrons from organic compounds such as glucose, maltose Obtains electrons from inorganic compounds such as H2, H2O, H2S, NH3, and S

i. Phototroph ii. Chemotroph iii. Mixotroph

Obtains energy from light (usually solar energy) Obtains energy from oxidation of chemical compounds Obtains energy from light and chemical reactions

Source of electrons

Source of energy

Note:  Further classification involves combinations of those listed, e.g., photoheterotrophs, chemoheterotrophs, photoautotrophs, and chemoautotrophs.

the form of phosphorylation, transacylation, condensation, or oxidation-reduction (redox) reactions. Phosphorylation reactions are used to initiate fueling processes since they produce substrate derivatives which cannot diffuse out of the cell (and energy is therefore not required to retain the substrate inside the cell). Phosphorylation of a substrate additionally places it in a more labile form for metabolism such that the activation energy of subsequent enzyme-catalyzed conversions is lowered and the specificity of enzymatic reactions is promoted [5]. A good illustration is the initial step of glycolysis, where glucose is phosphorylated to glucose-6-phosphate in the two coupled half-reactions shown below:

i. Nonspontaneous/endergonic half-reaction: Pi + Glucose ↔ Glucose-6-phosphate + H2O  ∆G´10 = +14 kJ/mole

(2.4)

ii. Spontaneous/exergonic half-reaction: ATP + H2O ↔ ADP + Pi   ∆G´20 = −31 kJ/mole

(2.5)

where Pi is an inorganic phosphate. The overall reaction is: ATP + Glucose →ADP + Glucose-6-phosphate ∆G´30 = ∆G´10 + ∆G´20 = −17 kJ/mole

(2.6)

Energy obtained from fueling reactions is efficiently captured in the chemical bonds of compounds such as nicotinamide adenine dinucleotide (NADH), nicotinamide adenine dinucleotide phosphate (NADPH) and adenosine triphosphate (ATP) via reversible redox and phosphorylation reactions. The reaction step in the “pay-off” phase of glycolysis in which glyceraldehyde-3-phosphate is converted to 3-phosphoglycerate is an example of a fueling reaction that supplies ATP through substrate-level phosphorylation in many organisms (refer to Section 2.6.4). The overall reaction is written as: Glyceraldehyde-3-phosphate + ADP + Pi + NAD ↔ 3-Phosphoglycerate + ATP + NADH + H+  ∆G´0 = −12.5 kJ/mole (2.7) Coupled reactions are very common in catabolic processes. Endergonic reactions are coupled to exergonic reactions such as ATP hydrolysis or NAD(P)H oxidation, and their free energy changes are

2-4

Cellular Metabolism

additive. By coupling reactions, energy from the exergonic reaction shifts the endergonic reaction away from equilibrium and toward product formation. Bioenergetics of catabolic and energy-producing pathways is discussed in greater detail in Section 2.6.5.

2.4  Products of Fueling Processes Fueling processes produce precursor metabolites, high-energy group-transfer bonds, and reducing power (primarily NADPH) which are used in anabolic processes. For example, the requirements for production of 1g of E. coli include approximately 13.4 mmole of precursor metabolites in the specific proportions needed by the cell (refer to Table 2.2), 41.2 mmole of high-energy phosphate bonds (denoted as ~P) and 18.5 mmole of NADPH [4]. The productivity of a fueling process depends on the nature and availability of substrates, intracellular and extracellular conditions, and the redox potentials of available electron donors and acceptors. Precursor metabolites, ATP, NADH, and NADPH are described below.

2.4.1  Precursor Metabolites The precursor metabolites obtained from fueling reactions serve as the major link between catabolic and anabolic processes. They are used in the synthesis of macromolecular subunits such as amino acids, lipids, enzymes, and nucleotides. They can alternatively be oxidized for ATP synthesis. Central metabolism produces 12 precursor metabolites that are essential to biosynthesis [6]. These central pathways include: Embden–Meyerhof–Parnas pathway (EMP, also known as glycolysis), Entner–Doudoroff pathway (ED), pentose phosphate pathway (PPP) and the tricarboxylic acid (TCA) cycle. More information on the central pathways is presented in Section 2.6.2. Table 2.2 lists 12 important precursor metabolites, their main sources and the required proportions for synthesis of 1 g of E. coli.

2.4.2  ATP During metabolism, high-energy intermediate compounds spontaneously transfer specific chemical groups, such as phosphoryl or acyl groups, to other compounds, and hence are said to have high “grouptransfer” potential. Hydrolysis of ATP and other high-energy compounds yields a strongly negative ∆G Table 2.2  Twelve Major Precursor Metabolites and the Primary Pathway Used for their Production Precursor Metabolite Glucose-6-phosphate Fructose-6-phosphate Triose phosphate 3-Phosphoglycerate Phosphoenolpyruvate Pyruvate Ribose-5-phosphate Erythrose-4-phosphate Acetyl-CoA α-Ketoglutarate Oxaloacetate Succinyl-CoAa

Metabolic Pathway

Amount Required for Biosynthesis of E. coli (µmol/g cell)

Glycolysis Glycolysis Glycolysis Glycolysis Glycolysis Glycolysis PPP PPP TCA cycle TCA cycle

205 71 129 1,496 519 2,833 898 361 3,748 1,079

TCA cycle TCA cycle

1,787 –

Source: Ingraham, J.L., Maaloe, O., and F.C. Neidhardt. Growth of the Bacterial Cell, Sinaeur Associates, Inc, Sunderland, MA, 1983. With permission. Note: Also included are the specific proportions of these metabolites required in the synthesis of E. coli. a Succinyl-CoA is a precursor metabolite in the synthesis of tetrapyrroles.

2-5

Catabolism and Metabolic Fueling Processes Table 2.3  Free Energy of Hydrolysis of Important High-Energy Compounds in Cellular Metabolism Compound

∆G´0 of Hydrolysis (kJ/mol)

Acetyl-CoA

−31.4

Phosphoenolpyruvate (PEP)

−61.9

Phosphocreatine

−43.0

Pyrophosphate

−19.2

ATP (to ADP)

−30.5

ATP (to AMP)

−45.6

ADP (to AMP)

−32.8

AMP (to Adenosine + Pi) Glucose-6-phosphate

−14.2

Fructose-6-phosphate

−15.9

Glycerol-1-phosphate

−9.2

−13.8

Source: Nelson D.L. and M.M. Cox, Lehninger Principles of Biochemistry, 4th ed., W.H. Freeman and Company, New York, 2005. With permission.

(usually more negative than −25 kJ/mole [5]), and is usually coupled to endergonic reactions. Table 2.3 lists the free energy of hydrolysis of some high-energy compounds. ATP (shown in Figure 2.1) is the most commonly used high-energy compound in the cell. The symbol “~” is used to denote high-energy bonds, hence ATP is also written as A ~ P ~ P ~ P. ATP is generated within the cell by phosphorylation of ADP through substrate-level phosphorylation and oxidative phosphorylation (refer to Sections 2.6.4 and 2.6.5). The intermediate ∆G for ATP hydrolysis along with its high phosphate group-transfer potential make it the universal energy currency in metabolic reactions. ATP hydrolysis involves the removal of one or two of its phosphates as shown:

ATP + H2O ↔ ADP + Pi + Energy   ∆G´0 = −30.5 kJ/mole

(2.8)



ADP + H2O ↔ AMP + Pi   ∆G´0 = −32.8 kJ/mole

(2.9)



ATP + H2O ↔ AMP + PPi   ∆G´0 = −45.6 kJ/mole

(2.10)



AMP + H2O ↔ Adenosine + Pi   ∆G´0 = −14.2 kJ/mole

(2.11)



PPi ↔ 2Pi   ∆G´0 = −19.2 kJ/mole

(2.12)

where Pi is a phosphate group and PPi is a pyrophosphate group. All values in Equations 2.8–2.12 were obtained from Ref. [5]. Structures of ATP, ADP, and AMP are shown in Figure 2.1. Hydrolysis of phosphoanhydride bonds in ATP is strongly exergonic because the resonance stabilization of hydrolysis products exceeds the resonance stabilization of the original compound. Also, the electrostatic repulsions between negatively charged phosphate oxygen atoms favor their separation. ATP hydrolysis is slow in the absence of an enzyme catalyst, and AMP and ADP levels regulate the hydrolysis of ATP in the cell (i.e., ATP hydrolysis is most favorable at low concentrations of ADP and AMP). The actual free energy change of ATP hydrolysis under physiological conditions is different from the standard free energy change due to the unequal cellular concentrations of ATP, ADP and Pi, their ionization states at cellular pH, and their formation of complexes with Mg2 + :

HATP3− ↔ ATP4− + H+  pK´ = 6.95 1

(2.13)



HADP2− ↔ ADP3− + H+  pK´ = 6.88 2

(2.14)

O–

P

O

O

OH

AMP

H

H

O

H OH

H

N

–O

O–

P

O

O

Inorganic phosphate

N

N

O

–

O–

P

O O O–

P

O O

OH

ADP

H

H

O H OH

H

N

N N

–O

N

NH2

O–

P

O O

O

–

O–

P

O O

O–

P

O O

O–

P

O O

OH

O

ATP

H

H

H OH

H

N

N N

N

NH2

Figure 2.1  Structures of adenosine monophosphate (AMP), adenosine diphosphate (ADP), and adenosine triphosphate (ATP). Adenosine is linked to phosphate through an ester bond. Other phosphate groups are linked by phosphoanhydride bonds.

–O

N

NH2

2-6 Cellular Metabolism

Catabolism and Metabolic Fueling Processes

2-7



H2PO4 − ↔ HPO42− + H+  pK´ = 7.20 3

(2.15)



Mg2+ + ATP4− ↔ MgATP2−

(2.16)



Mg2+ + ADP3− ↔ MgADP−

(2.17)



Mg2+ + HPO42− ↔ MgHPO4

(2.18)

Other high-energy compounds found in organisms include cytosine triphosphate (CTP), uridine triphosphate (UTP), guanosine triphosphate (GTP), phosphoenolpyruvate (PEP), acyl thioesters such as acetyl-coenzyme A (AcCoA), phosphocreatine in vertebrates, and phosphoarginine in invertebrates. ­High-energy compounds other than ATP, with higher or lower ∆Gs of hydrolysis and lower phosphate group-transfer potential, are used only in specific metabolic reactions. For example, CTP is used mainly in phospholipid synthesis, UTP is used for complex carbohydrate synthesis (such as glycogen and ­cellulose), and phosphocreatine is used mainly in skeletal muscle to provide chemical energy for intense work and to ensure a constant supply of ATP.

2.4.3 NADPH Nicotinamide adenine dinucleotide (NAD + , NADH) and nicotinamide adenine dinucleotide phosphate (NADP + , NADPH) belong to a class of molecules called mobile electron carriers (refer to Section 2.5) [5]. These water-soluble molecules migrate between enzymes and serve as coenzymes or enzyme cofactors in biological redox reactions. NADP + is similar to NAD + , with the addition of a phosphate group at the adenosine ribosyl C2 position. Reducing power extracted from substrates during oxidative fueling reactions is commonly stored in the nicotinamide ring of NAD(P)+ and the reduction is represented by the following reactions:

NAD +  + 2H +  + 2e − ↔ NADH + H+ ∆G'0 ≈ +61.9 kJ

(2.19)



NADP +  + 2H +  + 2e − ↔ NADPH+ H + ∆G'0 ≈ +62.5 kJ

(2.20)

Two hydrogen atoms are removed from the substrate, with one hydride ion (H: −) transferred to the nicotinamide ring and the other hydrogen released into solution as a free proton (H+ ). Figure 2.2 shows the structures of NAD(P)(H). NADP + reduction occurs most commonly through the oxidative pentose phosphate cycle and the TCA cycle, while NADH is generated from NAD + through glycolysis and the TCA cycle. In many organisms, reducing equivalents can be transferred between cofactors by the transhydrogenase (THD)catalyzed reaction:

NADPH + NAD +  ←transhydrogenase → NADP +  + NADH

(2.21)

The above reaction (Keq~1) is catalyzed by a cytosolic THD. Another membrane-bound, protontranslocating THD complex couples a proton gradient to the hydride transfer reaction, effectively shifting the equilibrium ratios of oxidized and reduced cofactors [7]:

[H + ]in + NADPH + NAD +  ←transhydrogenase → NADP +  + NADH + [H + ]out

(2.22)

Microbes often contain one of the two THD isoforms, and the membrane version is commonly found in mitochondria. Only the Enterobacteriaceae are known to contain both THDs. Information on the structural, catalytic and kinetic properties of transhydrogenases is available [7–10]. In prokaryotes

2-8

Cellular Metabolism H

O C

O –O

P

–O

P

O

O

O– H

O

2e– + H+

H

H

OH

H OH N

OH NAD+

–O

P

H

OH

H OH

OH

–O

P

H

O N

–O

P

H

O

–O

P

NH2

O

O–

H

H

OH

H OH

NH2 N

O–

N

N O

CH2 H

N

O

H

H

OH

O

H –O

O

O NADP+

H O

N

CH2

H O

H OH OH NADH

C

NH2

N

N

H

NH2

O

N

O

H

N

H

CH2

O

O–

NH2 N

O

2e– + H+

H

CH2 H

H OH

H

N O

OH

N

O H

O

H

O–

O

+ N

CH2 O–

H

H

C

P

H

H OH

H

–O

P

O

O–

H

H H

–O

N

O

P

NH2

N

CH2

O N

N

CH2

O

–O

NH2

O–

O

C

NH2

+ N

CH2

H O

H

O P

O

O NADPH

Figure 2.2  Structures of NAD(P)(H). Nicotinamide adenine dinucleotides serve as electron-shuttling coenzymes in metabolic reactions. NADP+ is similar to NAD+, with the addition of a phosphate group at the adenosine ribosyl C2 position.

(especially Enterobacteriaceae), the physiological role of transhydrogenases remains an intriguing area of study [11] and the importance of these enzymes in engineering cofactor-dependent metabolic pathways cannot be overstated (several noteworthy examples include Refs. [11–15]). Cofactor-dependent enzymes are typically specific for either NAD(H) or NADP(H), although some redox enzymes (e.g., many aldo-keto reductases) are capable of utilizing both cofactors. NAD + serves as an intermediate electron acceptor in catabolic reactions and as an electron shuttle in the respiratory chain. In contrast, NADPH typically serves as a hydride donor for polymerization and biosynthetic (anabolic) reactions such as fatty acid biosynthesis, steroid biosynthesis, and nucleic acid biosynthesis.

2-9

Catabolism and Metabolic Fueling Processes

NADPH additionally serves as a direct antioxidant that helps to protect eukaryotic cells from the damaging effects of oxygen radicals and thereby fight against diseases including atherosclerosis, cancer, and neurodegenerative disorders [5,16].

2.5  Redox Potentials and Mobile Electron Carriers 2.5.1  Redox Potentials The redox potential of a reaction is an important measure of its electron donating ability. Half-reactions involving molecules with low (usually negative) reduction potentials will donate electrons to those with high (positive) reduction potentials. In general, molecules with large negative reduction potentials are good electron donors and those with large positive reduction potentials are good electron acceptors. Table 2.4 lists some half-reactions occurring within cells, and their corresponding standard redox potentials. The relationship between the standard redox potential ( E ′ 0 ) and the actual redox potential (E) is:

E = E ′0 +

RT [Electron acceptor] ln [Electron donor] nF

(2.23)

where n is the number of electrons transferred per molecule, and F is the Faraday constant (F = 96,485 Coulombs/mole). For a redox reaction,

0 ΔE ′ 0 = E acceptor - E donor ′0 ′

(2.24)

Δ E ′ 0 is proportional to the free energy change ( ΔG ′ 0 ) as follows: ΔG ′ 0  = −nF∆ E ′ 0 (J)



(2.25)

An example calculation is given below, for the reduction of molecular oxygen (higher electrode potential) by FADH2 (lower electrode potential): Half-reactions: FAD + 2H +  + 2e −→FADH2 



E1′0  = −219 mV

(2.26)

1 O2 + 2H +  + 2e −→H2O  E ′20  = + 816mV 2

(2.27)

FADH2 + ½O2→FAD + H2O  ∆ E ′ 0  = + 1035mV

(2.28)

Overall reaction:

The ΔG ′ 0 of the overall reaction is:

ΔG ′ 0  = −2 × 96,485 × 1.035 ≈ −200 kJ

(2.29)

2.5.2  Mobile Electron Carriers Enzyme-catalyzed redox reactions are at the heart of catabolic processes, facilitating the conversion of substrate chemical energy to cellular energy currency. Hydrogen atoms and electrons obtained from redox reactions are transported in the cell by mobile electron carriers which include the flavoproteins, quinones, iron-sulfur proteins, cytochromes, and NAD(P)(H). The mobile electron carrier used in a particular redox reaction is dependent on the reduction potential and the chemical structure of the carrier.

2-10

Cellular Metabolism Table 2.4  Standard Redox Potentials of Some Cellular Reactions Couple Ferredoxin(Fe3 +)/ferredoxin(Fe2 +) (spinach) CO2/formate

E´0 (mV) −432 −432

H + /H2 (pH = 7) Ferredoxin (oxidized/reduced) in Clostridium

−410a

Acetoacetate/hydroxybutyrate

−346

−410

NADP + /NADPH

−324

NAD + /NADH FeS (oxidized/reduced) in mitochondria

−320 −305

Lipoic acid/dihydrolipoic acid

−290

S0/H2S

−270

FAD/FADH2

−220b

Acetaldehyde/ethanol

−197

FMN/FMNH2

−190

Pyruvate/lactate

−185

Oxaloacetate/malate

−170

Menaquinone (oxidized/reduced) Cyt b558 (oxidized/reduced)

−74 −75 to −43

Crotonyl-CoA/butyryl-CoA

−15

Fumarate/succinate

 + 33

Ubiquinone (oxidized/reduced)

 + 40 to + 100

Cytochrome b556 (oxidized/reduced)

 + 46 to + 129

Cytochrome b562 (oxidized/reduced)

 + 125 to + 260

Cytochrome d (oxidized/reduced)

 + 260 to + 280

Cytochrome c (oxidized/reduced)

 + 250

FeS (oxidized/reduced) in mitochondria

 + 280

Cytochrome a (oxidized/reduced)

 + 290

O2/H2O2

 + 295

Cytochrome c555 (oxidized/reduced)

 + 355

Cytochrome a3 (oxidized/reduced) in mitochondria

 + 385

NO3−/NO2−

 + 421

Fe3 + /Fe2 +  O2 (1 atm)/H2O

 + 815

 + 771

Source: White, D. “Standard Electrode Potentials” from The Physiology and Biochemistry of Prokaryotes, Oxford University Press, New York, table 4.1, pg. 104, 2000. With permission. Some data also obtained from Nelson, D. and Cox, M. Lehninger Principles of Biochemistry. W.H. Freeman and Company, New York, 2005. Note: In developing the values for the standard redox potentials, a hydrogen electrode is arbitrarily designated as the reference, with [H+ ] = 1 M. The standard redox potential (E′ 0 ) is based on assumptions of 1M of electron acceptor and electron donor, pH = 7 and standard conditions of temperature and pressure (298 K and 1 atm). a When this half reaction occurs at pH = 0, E´0 is 0.0. b This reaction has different E´0 depending on whether FAD is free or if it is bound tightly to a flavoprotein. The value given is for free FAD.

2.5.3  Quinones Quinones serve as electron acceptors in the electron transport chains (ETCs) of photosynthetic photosystems I and II, aerobic and anaerobic respiration [6]. These hydrophobic, lipid-soluble electron carriers transport hydrogen and electrons to and from membrane-bound protein complexes. Three types of quinones are commonly found in ETCs:

2-11

Catabolism and Metabolic Fueling Processes CH3

O O

H

H3C

n

H3C CH3

O

Ubiquinone

O H+ + e–

O·

CH3 H

O H 3C

n

H3C CH3

O

Ubisemiquinone radical

OH H+ + e–

CH3

OH O

H

H3C

n

H3C O

CH3

Ubiquinol

OH

Figure 2.3  Reduction of ubiquinone. Reduction of ubiquinone occurs in two electron transfers through a ubisemiquinone radical intermediate. The value n can vary from 4 to 10.



1. Ubiquinone (coenzyme Q/CoQ): found in the mitochondrial and bacterial ETCs and characterized by two methoxy groups, as shown in Figure 2.3. 2. Menaquinone: found in the bacterial electron transport chain (ETC). It is characterized by a benzene ring replacing the two methoxy groups present in ubiquinones. Menaquinones have a lower electrode potential ( E ′ 0  = −74 mV) than ubiquinones ( E ′ 0  = +40 to +100mV) and are more commonly used during anaerobic respiration [6].

2-12



Cellular Metabolism

3. Plastoquinone: found in the ETCs of photosynthetic organisms such as plants and cyanobacteria. It is characterized by methyl groups which replace the two methoxy groups found in ubiquinone.

Figure 2.3 shows the reduction of ubiquinone to ubiquinol. An example of a quinone-mediated electron shuttling process is shown below:

CoQH2 + 2Fe3+ –cytochrome c→CoQ + 2Fe2+ –cytochrome c + 2H+ 

(2.30)

2.5.4  Flavoproteins Flavoproteins serve as enzyme catalysts in the redox reactions of oxidative phosphorylation and photophosphorylation. They employ either flavin mononucleotide (FMN or riboflavin-5′-phosphate) or flavin adenine dinucleotide (FAD) as their coenzyme and electron carrier. These coenzymes are depicted in Figure 2.4. The flavin nucleotides are tightly bound or in some cases covalently bound prosthetic groups of the flavoproteins. FAD and FMN can accept one or two electrons, with the fully reduced forms written as FADH2 and FMNH2.

2.5.5  Iron-Sulfur Proteins Iron-sulfur proteins (also called ferredoxins) are a group of electron carriers containing a 1:1 ratio of sulfide ions and nonheme iron ions which exist in a variety of oxidation states. The iron-sulfur proteins cover a very wide range of potentials, from approximately −400 to +350 mV; therefore, they are involved in redox reactions at the low and high ends of the potential range in the ETC [6]. The low potential ferredoxins can be either Fe2S2 ferredoxins or Fe4S4 ferredoxins [17–19] and they are found in plants, mitochondrial and bacterial electron transport systems. Specific examples include adrenodoxin in mitochondrial monooxygenase systems [20], thioredoxin-type ferredoxins in Clostridium pasteurianum [21,22] and putidaredoxin and terpredoxin in other bacterial systems. The high potential ferredoxins (HiPIP) are mainly Fe4S4 ferredoxins [17–19] found in anaerobic ETCs, and are common in photosynthetic bacteria [23]. High potential Fe4S4 ferredoxins have higher oxidation states than low potential Fe4S4 ferredoxins and are reduced as follows:

[Fe4S4 ]3+

High potential ferredoxin

←e →

[Fe4S4 ]2+

←e →

[Fe4S4 ]+

Low potential ferredoxin

(2.31)

2.5.6  Cytochromes Cytochrome proteins catalyze redox reactions in the ETC. They are ubiquitous in nature, found in plants, photosynthetic microorganisms, bacteria, the mitochondrial inner membrane and the endoplasmic reticulum of eukaryotes. They have an iron-containing heme prosthetic group which consists of four porphyrins (pyrrole rings with substituted side chains) attached by methane bridges, and this heme group is tightly bound or covalently linked to its associated protein [5]. Cytochromes are single electron carriers, and the iron ion in the heme transitions between Fe2+ and 3+  Fe states during the electron-transfer process. There are five different classes of hemes that distinguish the cytochromes: hemes a, b, c, d, and o. A cytochrome can have more than one heme group. For example, bacterial cytochrome bd contains hemes b and d, and it is also called cytochrome d. Similarly, bacterial cytochrome bo contains hemes b and o, and it is also called cytochrome o or cytochrome bo3 (with the subscript 3 indicating that it is an O2-binding heme). Hemes d and o are found only in the prokaryotic cytochrome oxidases [6,24].

H

O–

P

O

NH

O

O

FAD

O–

P

O O H OH

H

O H OH

H

N

N N

NH2 N

e–+ H+

CH3

CH3

H

H

H

FMNH·

H

H

CH2

OH

OH

OH

·C

CH2

N

H N

O

N

O

O

O–

P

O

NH

O–

e–+ H+

CH2

CH 2

H

H

H

H

H

O

OH

OH

OH

N H

O

FMNH2

CH2

CH2

N

H N

O

O–

P

O

NH

O–

Figure 2.4  Reduction of FMN and FAD. Flavin mononucleotide (FMN) and flavin adenine dinucleotide (FAD) are derivatives of riboflavin. Phosphorylation of riboflavin at the ribityl 5´-OH produces FMN and adenylation of FMN produces FAD. Their reduction chemistry is illustrated, using FMN as an example. Complete reduction of FMN and FAD requires a two-step electron transfer.

FMN

OH

H

O

OH

OH

N

O

H

CH2

CH2

N

CH3

H

N

CH3

H

Catabolism and Metabolic Fueling Processes 2-13

2-14

Cellular Metabolism

2.6 Examples of Catabolic Processes in Different Organisms 2.6.1  Photosynthesis Phototrophic organisms use light to generate chemical free energy, which is stored in the bonds of ATP and NAD(P)H in the catabolic phase of photosynthesis. In the anabolic phase of photosynthesis, ATP and NADPH are utilized in the synthesis of carbohydrates and other organic compounds from CO2 and H2O. Examples of phototrophic organisms include green plants, algae, and photosynthetic bacteria. These organisms carry out either oxygenic (O2-producing) or anoxygenic (non-O2producing) photosynthesis. 2.6.1.1 Oxygenic Photosynthesis Oxygenic photosynthesis is carried out by green plants, algae, the cyanobacteria and photosynthetic bacteria of the genera Prochloron, Prochlorothrix and Prochlorococcus [6]. Oxygenic photosynthesis involves a “light phase” and a “dark phase.” The light phase is necessary in oxygenic photosynthesis because H2O, an inherently poor electron donor is used, and the transfer of electrons from H2O (converted to O2, ∆E′ 0  = +820 mV) to the reaction center chlorophyll, P680 (∆E′ 0 = +1.1 mV) is driven by light energy [6]. The main light-absorbing pigment found in green plants and cyanobacteria is chlorophyll a, and in the prochlorophytes it is chlorophyll b. The light-dependent reactions generate NADPH and a proton gradient (∆p) which is dissipated for ATP synthesis (discussed in Section 2.6.5). Two light reactions occur at two different reaction centers: Reaction center I (Photosystem (PS) I), which is energized by light at wavelengths of about 700 nm or higher, and reaction center II (Photosystem (PS) II), which is energized by light at wavelengths of about 680 nm. The activities of PS I and II are coupled and are connected by a cyclic or noncyclic ETC through which electrons are transferred from the final electron acceptor in PS II to the PS I chlorophyll, with the concomitant generation of a proton gradient. For every mole of O2 formed, 12 protons move across the thylakoid membrane and about 200 kJ of energy is stored in the resulting proton gradient. This energy is used to synthesize about three moles of ATP [5]. The reactions of the light phase are summarized below:

2H2O + 2NADP+  + eight photons + ~3ADP + ~3Pi → O2 + 2NADPH + ~3ATP

(2.32)

2.6.1.2  Anoxygenic Photosynthesis Anoxygenic photosynthesis is carried out by most photosynthetic bacteria. The major classes of anoxygenic phototrophs are Rhodospirillaceae (purple nonsulfur bacteria), Chromatiaceae (purple sulfur bacteria), Chlorobiaceae (green sulfur bacteria), Chloroflexaceae (green nonsulfur bacteria/green gliding bacteria), and heliobacteria [6,25–27]. The main light-absorbing pigment in anoxygenic phototrophs is the bacterio-chlorophyll. There are seven types of bacterio-cholorophyll and they are distinguished by their respective absorption spectra [28]. Inorganic sources of electrons such as H2S, S0, S2O32− , and H2 are utilized, and the oxidation of these compounds is not necessarily light-dependent. Only one reaction center is necessary for anoxygenic photosynthesis, and electron flow can be cyclic or noncyclic [6,26,28]. The photosynthetic mechanisms in the purple bacteria are similar to those of PS II in oxygenic photosynthesis, while photosynthesis in green bacteria and heliobacteria resembles that of PS I [6]. Below is an example of a light phase reaction during anoxygenic photosynthesis in green sulfur bacteria [6]:

+ bacteriochlorophyll S 0 + NADH + H +  + ATP H2S + NAD +  + ADP + Pi light  →

(2.33)

Products of oxygenic and anoxygenic photosynthesis light phase reactions are employed in the dark phase for the reduction of CO2 to triose phosphate in a process called carbon fixation. Carbon fixation commonly occurs through the Calvin cycle. In plants and algae, Calvin cycle reactions take place in the stroma of chloroplasts, while in photosynthetic bacteria they take place in the cytosol [6]. For every

Catabolism and Metabolic Fueling Processes

2-15

mole of triose phosphate produced, three moles of CO2, six moles of NADPH, and nine moles of ATP are utilized. The triose phosphates are further converted to starch and hexoses in the stroma and cytosol, respectively. Carbon fixation is also accomplished through pathways other than the Calvin cycle. A reductive or reverse TCA cycle is used to fix CO2 by Chlorobiaceae (a class of green sulfur bacteria), Desulfobacter (anaerobic sulfate-reducing bacterium), Hydrogenobacter, and archaea such as Thermoproteus neutrophilus (note that additional enzymes are required for a functional, reverse TCA cycle) [6,29–31]. The 3-hydroxypropionate cycle is a recently discovered carbon fixation pathway demonstrated in bacteria and archaea such as Chloroflexus aurantiacus (a green nonsulfur bacteria) and Metallosphaera sedula (a hyperthermophilic crenarchaeon) [31,32]. Finally, a noncyclic acetyl-CoA pathway fixes CO2 in methanogens, acetogenic bacteria, and most autotrophic sulfate-reducing bacteria [33].

2.6.2 The Central Pathways The central pathways are used in the metabolism of carbohydrates and carboxylic acids. Central pathways produce precursor metabolites (carbon skeleton) and sometimes ATP and NAD(P)H (energy), in which case they are called amphibolic pathways. Examples of amphibolic pathways are: the EMP pathway (glycolysis), the ED pathway, the PPP, and the TCA cycle (shown in Figures 2.5 and 2.6). The EMP, PPP and ED pathways are generally considered in the context of glucose oxidation to pyruvate, with each pathway employing different enzymes/reactions to synthesize the intermediate glyceraldehyde-3phosphate. However, in some bacteria including E. coli, the ED pathway is used primarily for gluconate metabolism rather than glucose metabolism [4]. 2.6.2.1  Glycolysis Glycolysis comprises the initial reactions required for carbohydrate metabolism. The principal functions of glycolysis are: oxidation of glucose (a six carbon molecule) to pyruvate (a three carbon molecule), production of cellular energy sources (NADH and ATP), and supply of six of the precursor metabolites used in biosynthesis (refer to Table 2.2). The reactions of glycolysis are summarized as: Glucose + 2NAD +  + 2ADP + 2Pi→2Pyruvate + 2NADH + 2H +  + 2ATP + 2H2O ∆G′ 0  = −85 kJ/mole

(2.34)

Glucose oxidation by glycolysis is incomplete and the final product (two pyruvates) still contains the bulk of the total energy initially in glucose. Complete oxidation of one mole of glucose yields about 2,840 kJ of energy, but 94.8% of this energy is contained in two moles of pyruvate [5]. Over 60% of the energy released from glycolysis is stored as ATP [5]. During glycolysis, ATP is produced only by substrate-level phosphorylation, whereby energy released from exergonic reactions is used to drive the phosphorylation of ADP [25]. For example:

pyruvate

kinase 2Phosphoenolpyruvate + 2ADP  → 2Pyruvate + 2ATP  ∆G′ 0  = −31.4 kJ/mole

(2.35)

Glycolysis is divided into two stages: the preparatory phase and the pay-off phase. The preparatory phase consists of the first five steps which incorporate isomerization, group-transfer and cleavage reactions. Two moles of ATP are utilized and two moles of glyceraldehyde-3-phosphate are produced per mole of glucose. The pay-off phase is comprised of redox reactions which oxidize glyceraldehyde-3phosphate to pyruvate. The reactions in this phase yield four moles of ATP and two moles of NADH per mole of glucose. The reactions involved in glycolysis are summarized in Table 2.5. Note that several of these reactions are considered irreversible.

2-16

Cellular Metabolism ED

EMP

PPP

Glucose + ATP NADP+ NADPH NADPH NADP 1 NADP+ NADPH ADP Ribulose-5-phosphate Glucose-6-phosphate 6-Phosphogluconate 6-Phosphogluconate 11 11 12 14 2 13 CO 2 18 Fructose-6-phosphate Ribose-5-phosphate Xylulose-5-phosphate 2-Keto-3-deoxy-6ATP 3 phosphogluconate 15 ADP Fructose-1,6-bisphosphate SedoheptuloseGlyceraldehyde4 7-phosphate 3-phosphate Glyceraldehyde-3-phosphate 19 16 + Dihydroxyacetonephosphate FructoseErythrose6-phosphate 5 4-phosphate Glyceraldehyde-3-phosphate NAD+ + Pi 6 NADH 1,3-Bisphosphoglycerate ADP 7 ATP 3-Phosphoglycerate

17 Xylulose5-phosphate

8 2-Phosphoglycerate 9 Phosphoenolpyruvate ADP 10 ATP Pyruvate

Fermentation products

TCA cycle/anaplerosis Biosynthesis

Figure 2.5  The central pathways. The glycolysis/Embden-Meyerhof-Parnas (EMP) pathway converts glucose6-phosphate to pyruvate. The PPP and the ED pathway branch from 6-phosphogluconate and re-enter glycolysis at different locations. In the PPP, the oxidation-decarboxylation reactions producing ribulose-5-phosphate occur in four steps: oxidation of glucose-6-phosphate to 6-phosphogluconolactone, hydrolysis of 6-phosphogluconolactone to 6-phosphogluconate, oxidation of 6-phosphogluconate to 3-keto-6-phosphogluconate and decarboxylation of 3-keto-6-phosphogluconate. These reactions form the oxidative arm of the PPP. The remaining reactions form the reductive arm of the PPP and consist of isomerizations and sugar rearrangements. The ED pathway produces one mole of glyceraldehyde-3-phosphate per mole of glucose-6-phosphate, so the net ATP yield up to pyruvate synthesis is only one in this pathway (instead of two, as in EMP). The enzymes that catalyze the reactions are: 1, hexokinase; 2, phosphohexose isomerase; 3, phosphofructokinase-1; 4, aldolase; 5, triose phosphate isomerase; 6, glyceraldehyde-3-phosphate dehydrogenase; 7, phosphoglycerate kinase; 8, phosphoglycerate mutase; 9, enolase; 10, pyruvate kinase; 11, glucose-6-phosphate dehydrogenase + gluconolactonase; 12, 6-phosphogluconate dehydrogenase; 13, ribose-5-phosphate isomerase, 14, ribulose-5-phosphate epimerase; 15, transketolase; 16, transaldolase; 17, transketolase; 18, 6-phosphogluconate dehydratase; 19, 2-keto-3-deoxy-6-phosphogluconate aldolase.

2.6.2.2 The PPP The PPP, also called the phosphogluconate pathway or the hexose monophosphate pathway, is responsible for the oxidation of glucose-6-phosphate to pentose phosphates such as ribose-5-phosphate and xylulose-5-phosphate. These pentose phosphates are used in the synthesis of RNA, DNA, and coenzymes such as ATP, coenzyme A, NADH, and FADH2. Additionally, the PPP produces glycolytic intermediates by converting five-carbon sugars (more accurately their phosphates) to six-carbon and three-carbon sugars, and it also produces two of the precursor metabolites used in biosynthesis (refer to Table 2.2).

2-17

Catabolism and Metabolic Fueling Processes H2O

FADH2

FAD

Fumarate 7 Malate

NAD+

Succinate Acetyl-CoA

10

CoA

ADP/GDP + Pi 9

Succinyl-CoA NADH + CO2 4

Citrate

H2O

CoA + ATP/GTP

5

Glyoxylate

Oxaloacetate

1

H2O

CoA

8

NADH

Acetyl-CoA

6

2a

NAD+ + CoA

cis-Aconitase

α-Ketoglutarate 2b

Isocitrate

3

NAD(P)H + CO2

NAD(P)+

H2O

Figure 2.6  The tricarboxylic acid (TCA/citric acid/Krebs) cycle. Acetyl-CoA is oxidized to CO2 and energy is conserved as NAD(P)H, FADH 2 and ATP. Enzymes involved: 1, citrate synthase; 2a, aconitase; 2b, aconitase; 3, isocitrate dehydrogenase; 4, α-ketoglutarate dehydrogenase complex; 5, succinyl-CoA synthetase; 6, succinate dehydrogenase (in oxidative direction) or fumarate reductase (in reductive direction); 7, fumarase; 8, malate dehydrogenase; 9, isocitrate lyase; 10, malate synthase. In eukaryotes, the enzymes of the citric acid cycle are found in the mitochondrial matrix while in prokaryotes they are cytosolic (succinate dehydrogenase interacts with the membrane). Depending on the isoform, isocitrate dehydrogenase requires either NADP+ or NAD+. Different isoforms of succinyl-CoA synthetase generate either ATP or GTP through substrate-level phosphorylation.

Table 2.5  Reactions of Glycolysis and Their Standard Free Energies ∆G´0 (kJ/mole) A. Preparatory Phase   (1)

Glucose

—▶

Glucose-6-phosphate

  (2)

Glucose-6-phosphate

◀—▶

Fructose-6-phosphate

  (3)

Fructose-6-phosphate

—▶

Fructose-1,6-bisphosphate

  (4)

Fructose-1,6-bisphosphate

◀—▶

Glyceraldehyde-3-phosphate  + Dihydroxyacetone phosphate

  (5)

Dihydroxyacetone phosphate

◀—▶

Glyceraldehyde-3-phosphate

−16.7 1.7 −14.2 23.8 7.5

B. Pay-off Phase   (6)

Glyceraldehyde-3-phosphate

◀—▶

1,3-Bisphosphoglycerate

  (7)

1,3-Bisphosphoglycerate

◀—▶

3-Phosphoglycerate

6.3 −18.5

  (8)

3-Phosphoglycerate

◀—▶

2-Phosphoglycerate

4.4

  (9)

2-Phosphoglycerate

Phosphoenolpyruvate

7.5

(10)

Phosphoenolpyruvate

◀—▶ —▶

Pyruvate

−31.4

Source: Nelson, D. and Cox, M. Lehninger Principles of Biochemistry. W.H. Freeman and Company, New York, 2005. With permission.

2-18

Cellular Metabolism

The PPP is the main source of NADPH, which serves as an electron donor in most biosynthesis reactions. The reactions of the PPP are divided into one oxidative phase and two reductive phases [6]. Several of the reactions in the reductive phase of the PPP are similar to those used by autotrophic organisms to fix CO2 via the Calvin cycle. Stage 1: Oxidation–decarboxylation reactions

+

NADP →6 NADPH → 3Ribulose-5-phosphate + 3CO2 3Glucose-6-phosphate + 3H2O 6

(2.36)

Stage 2: The isomerization reactions

3Ribulose-5-phosphate → 2Xylulose-5-phosphate + Ribose-5-phosphate

(2.37)

Stage 3: The sugar rearrangement reactions

i. Transketolase-catalyzed reaction Xylulose-5-phosphate + Ribose-5-phosphate → Sedoheptulose-7-phosphate + Glyceraldehyde-3-phosphate

(2.38)

ii. Transaldolase-catalyzed reaction Sedoheptulose-7-phosphate + Glyceraldehyde-3-phosphate → Fructose-6-phosphate + Erythrose-4-phosphate

(2.39)

iii. Transketolase-catalyzed reaction

Xylulose-5-phosphate + Erythrose-4-phosphate → Fructose-6-phosphate + Glyceradehyde-3-phosphate

(2.40)

Two moles of fructose-6-phosphate produced by reactions 2.39 and 2.40 can be isomerized back to two moles glucose-6-phosphate. The net reaction of the PPP is thus:

Glucose-6-phosphate + 6NADP +  + 3H2O → Glyceradehyde-3-phosphate + 3CO2 + 6NADPH + 6H + 

(2.41)

2.6.2.3 The ED Pathway The ED pathway is primarily found in prokaryotes but has been reported in Entamoeba histolytica and the fungi Aspergillus niger and Penicillum notatum [6,34]. In the ED pathway, one mole of glucose-6-phosphate is oxidized to one mole of glyceraldehyde-3-phosphate and one mole of pyruvate. Glyceraldehyde-3-phosphate is further oxidized through the EMP pathway to produce another mole of pyruvate. The ED pathway is summarized as:

Glucose + NADP +  + NAD +  + ADP + Pi → 2Pyruvate + NADPH + 2H +  + NADH + ATP

(2.42)

When the ED pathway is used for glucose oxidation, the net ATP yield per mole of glucose is one. Based on the lower energy yield of the ED pathway, anaerobes which lack respiratory chains use the EMP pathway (net ATP yield per mole of glucose is two) for glucose oxidation since this is more “economical” [25]. However the ED pathway is useful in those organisms that do not have a complete EMP pathway and also for the degradation of aldonic acids such as gluconate [6].

Catabolism and Metabolic Fueling Processes

2-19

The distribution of EMP and ED pathways in various bacteria is summarized by White [6, chapter 8]. Flux ratios through these “upper” metabolic pathways vary considerably between organisms and growth conditions. Metabolic engineers are often concerned with these fluxes and altering them through genetic modifications. Continued advances in flux analysis are enabling novel insights into cellular metabolism and improved strain engineering (for examples of flux analysis methods and applications refer to Refs [35–42], as well as section IV of this handbook). 2.6.2.4  Fates of Pyruvate Pyruvate is obtained from the breakdown of nearly all energy sources and its metabolic fate is highly dependent on growth conditions. Pyruvate can be siphoned-off for use as a precursor metabolite in the synthesis of the amino acids alanine, valine, and leucine [5] and it can be utilized by anaplerotic reactions (discussed in Section 2.6.3) involved in replenishing citric acid cycle intermediates. Alternatively, pyruvate can be converted into the high-energy intermediate acetyl-CoA, which plays an important role in connecting glycolysis to the citric acid cycle. Depending on the organism, conversion of pyruvate to acetyl-CoA can be catalyzed by the pyruvate dehydrogenase enzyme complex (PDHc), pyruvate formate lyase (PFL), or pyruvate:ferredoxin oxidoreductase (POR) and fluxes through these enzymes depend on whether metabolism proceeds via the oxidative route (citric acid cycle and respiration) or fermentative route [43]. PDHc catalyzes the NAD + -linked oxidative decarboxylation of pyruvate to acetyl-CoA (primarily considered an aerobic reaction) and PFL catalyzes the anaerobic conversion of pyruvate to acetyl-CoA [5,44–48], as shown: PDHc



→ Acetyl-CoA + NADH + H +  + CO2 Pyruvate + CoA + NAD +  

(2.43)



Pyruvate + CoA PFL → Formate + Acetyl-CoA

(2.44)

Thus, when cells are able to generate energy through respiration, pyruvate oxidation provides NADH and fully oxidized CO2, while under fermentative conditions NADH production is avoided by the production of formate. In E. coli, PDHc is generally considered to be minimally active under anaerobic or at least non-respiratory conditions [43] due to a relatively high ratio of NADH/NAD + and high concentration of acetyl-CoA, both of which allosterically inhibit PDHc activity [43,49,50]. However, when nitrate serves as an anaerobic terminal electron acceptor, PDHc activity in E. coli can be substantial [43,48]. Expression of PDHc is additionally repressed by the ArcA/B global regulator [51] under low oxygen conditions (or more accurately, low oxidation-reduction potentials) [47,51–54]. While PDHc activity is extremely redox sensitive in many organisms (especially Enterobacteriaceae), pyruvate dehydrogenases from some organisms such as Enterococcus faecalis and Azotobacter vinelandii are less sensitive to elevated levels of intracellular NADH [50,55]. PFL is enzymatically interconverted between active and inactive forms during anaerobic-aerobic transitions since its activity involves a glycyl radical that is not stable in the presence of oxygen [43,46,48]. At the genetic level, expression of PFL in E. coli is regulated by the FNR and ArcA/B global regulators [6,43,51]. In general, catabolic pathways are sensitive to and tightly regulated by a number of related signals that reflect the redox potential and energy charge of the cell (refer to Section 2.6.6 for details on regulation of catabolic pathways). Pyruvate can alternatively be converted directly to acetate and CO2 by pyruvate oxidase under aerobic conditions, in which case reducing equivalents directly enter electron transport through ubiquinone. Acetate can in turn be converted to acetyl-CoA through the ATP-dependent acetokinase-phosphotransacetylase (ACK-PTA) and acetyl-CoA synthetase (ACS) pathways [56,57].

Acetate + ATP ←ACK  → Acetyl-P + ADP

(2.45)

2-20

Cellular Metabolism



Acetyl-P + CoA ←PTA → Acetyl-CoA + Pi

(2.46)



CoA + Acetate + ATP ←ACS → Acetyl-CoA + AMP + PPi

(2.47)

Another fate of pyruvate is nonoxidative decarboxylation to acetaldehyde by the fermentative enzyme pyruvate decarboxylase (PDC). PDC from Zymomonas mobilis has received much attention in the metabolic engineering of fermentation pathways for ethanol production [58,59]. 2.6.2.5 The TCA Cycle The TCA/citric acid cycle is used in the metabolism of sugars, fatty acids, and some amino acids. Its reactions oxidize acetyl-CoA to CO2, and reducing equivalents are extracted in the form of NAD(P)H and FADH2. One mole of ATP is also produced per mole of acetyl-CoA through substrate-level phosphorylation. The citric acid cycle operates concurrently with the respiratory chain, through which NAD(P)H and FADH2 are reoxidized and a proton motive force is generated for ATP synthesis [6] (refer to Section 2.6.5). The overall reactions of the TCA cycle can be summarized as:

Acetyl-COA + 2H2O + ADP + Pi + FAD + NADP+  + 2NAD+ →2CO2 + ATP + FADH2  + NADPH + 2NADH + 3H+  + CoA

(2.48)

Although typically associated with aerobic respiration, the citric acid cycle can be active during anaerobic respiration in the presence of an external electron acceptor [60]. For example, Pseudomonas stutzeri completely oxidizes glucose to CO2 with nitrate as the electron acceptor [61,62] and the hyperthermophilic archaea, Thermoproteus tenax and Pyrobaculum islandicum, express a functional TCA cycle with sulfur or thiosulfate as the terminal electron acceptor [6,63]. Regardless of whether the TCA cycle is operational, enzymes of the cycle are used to generate precursor metabolites for biosynthesis [64] (refer to Table 2.2). For example, succinyl-CoA is a precursor to the synthesis of L-lysine, L-methionine, and tetrapyrroles of cytochromes and chlorophylls [6]. Under fermentative conditions the TCA cycle enzyme α-ketoglutarate dehydrogenase has little or no activity [51,64–66] and therefore cannot synthesize sufficient succinyl-CoA. For continued supply of metabolic precursors the TCA cycle enzymes then operate in two separate branches. α-ketoglutarate is supplied from citrate through the oxidative branch (as in the normal cycle), while succinyl-CoA is produced via reduction of oxaloacetate through the reverse, “reductive” branch, in which the enzyme fumarate reductase replaces succinate dehydrogenase.

2.6.3  Anaplerotic and Peripheral Reactions A variety of important reactions facilitate the operations of catabolic and anabolic pathways by connecting and replenishing their intermediates. Peripheral reactions convert substrates (carbon sources) which are non-central pathway intermediates into central pathway intermediates so that they can be subsequently metabolized. These reactions enhance the metabolic flexibility of organisms by enabling them to grow on a broader range of carbon compounds. For example, glycerol is converted to dihydroxyacetone phosphate (a glycolytic intermediate) in the two-step feeder pathway involving the enzymes glycerol kinase and glycerol-3-phosphate dehydrogenase. Anaplerotic reactions are responsible for replenishing central pathway intermediates which have been depleted through biosynthesis, and are most commonly referred to in the context of restoring TCA cycle intermediates. In a similar context, cataplerotic reactions refer to those which deplete excess TCA cycle intermediates, and play an important role in gluconeogenesis [67]. Anaplerosis derives carbons from amino acids, carboxylic acids, lipids, and CO2. The conversion of pyruvate to oxaloacetate by the

Catabolism and Metabolic Fueling Processes

2-21

pyruvate carboxylase enzyme is an example of an anaplerotic reaction which replenishes a citric acid cycle intermediate:

pyruvate

Pyruvate + HCO3− + ATP ←carboxylase  → Oxaloacetate + ADP + Pi

(2.49)

When oxaloacetate nears depletion and the citric acid cycle is unable to effectively process acetylCoA, excess acetyl-CoA accumulates in the cell and acts as a positive allosteric modulator of the pyruvate carboxylase enzyme, stimulating oxaloacetate production [5]. Oxaloacetate is also produced in the anaplerotic reaction catalyzed by PEP carboxylase:

PEP carboxylase

→ Oxaloacetate + Pi PEP + HCO3− ←

(2.50)

Depending on the organism, PEP carboxylase is regulated by metabolic intermediates such as acetyl-CoA [6] or fructose-1,6-bisphosphate [5]. A similar reaction is catalyzed by the enzyme PEP carboxykinase, although its primary physiological role is in the reverse direction, i.e., PEP synthesis from oxaloacetate:

PEP

Oxaloacetate + ATP/GTP ←carboxykinase  → PEP + CO2 + ADP/GDP

(2.51)

Nonglycolytic PEP is also synthesized by the enzymes PEP synthetase and pyruvate-phosphate dikinase, both using pyruvate as substrate and requiring hydrolysis of ATP to AMP:

synthetase Pyruvate + H2O + ATP PEP  → PEP + AMP + Pi



dikinase Pyruvate + ATP + Pi  → PEP + AMP + PPi

Pyruvate-phosphate

(2.52) (2.53)

Another important anaplerotic reaction is the synthesis of pyruvate from malate by the malic enzyme:

Malic enzyme

→ Pyruvate + NAD(P)H + CO2 L-malate + NAD(P)+  ←

(2.54)

This is a critical reaction to support growth on TCA cycle intermediates, and the reverse reaction has been used to improve succinate production by E. coli [68]. Anaplerotic reactions play an important role in the overproduction of metabolites derived from TCA cycle intermediates and are a common target in metabolic engineering (refer to Refs. [69–78]). 2.6.3.1 The Glyoxylate Bypass In some aerobic organisms, acetyl-CoA can be oxidized to succinate or other four-carbon intermediates of the citric acid cycle through the glyoxylate bypass. The glyoxylate bypass is an anaplerotic pathway [79] in which the two decarboxylation steps of the citric acid cycle are bypassed [6,64] and isocitrate is converted to succinate and glyoxylate by the enzyme isocitrate lyase. Glyoxylate condenses with acetyl-CoA to form malate in a reaction catalyzed by malate synthase, and malate undergoes a NAD + -dependent oxidation to form oxaloacetate. The reactions are summarized as:

2Acetyl-CoA + NAD + + 2H2O → Succinate + 2CoA + NADH + H+ 

(2.55)

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Cellular Metabolism

The glyoxylate cycle is present in plants, some microorganisms such as E. coli and some invertebrates, allowing these organisms to grow on acetate and convert fatty acids into metabolic intermediates [5,64,79]. Upregulation of the glyoxylate bypass in E. coli was shown to reduce overflow metabolism/ acetate secretion (refer to Section 2.6.6) [80] and has been implemented for the overproduction of succinate [81].

2.6.4  Fermentation Cell fermentation requires no external electron acceptors and no respiratory pathways. Instead, internal metabolites such as pyruvate and acetyl-CoA serve as substrates of reduction reactions, allowing for NAD + regeneration with concomitant secretion of (reduced) acids and alcohols (e.g., lactate and ethanol). Fermentation is a poor energy-yielding process with little assimilation of substrate carbon into biomass. The driving force for fermentation is glycolytic ATP production through substrate-level phosphorylation, which can only continue when coupled to NADH oxidation. Additional ATP is sometimes produced in the subsequent fermentative pathways (e.g., during acetate production in mixed acid fermentation). Fermentation products vary depending on the organism and degree of reduction of the growth substrate. For example, lactic acid bacteria such as Lactobacillus plantarum produce lactate, yeasts and some fungi such as Trichocladium canadense [82] produce ethanol, and members of Clostridrium produce butyrate, butanol, and acetone [6]. E. coli and other Enterobacteriaceae carry-out mixed-acid fermentation [28], secreting mixtures of lactate, ethanol, acetate, succinate and formate. Recently, E. coli has been the target of many metabolic engineering efforts aimed at generation of strains which exclusively overproduce a single fermentation product (e.g., [81,83–89]. Figure 2.7 shows common fermentation pathways found in different organisms.

2.6.5 Oxidative Phosphorylation The central concept in oxidative phosphorylation is the chemiosmotic theory, developed by a biochemist, Peter Mitchell [90]. Cells use ion gradients to couple exergonic reactions which occur during electron transport to endergonic reactions such as ATP synthesis. During oxidative phosphorylation, electrons are ferried from NADH/FADH2 to terminal electron acceptors through protein complexes and mobile electron carriers in the ETC. Free energy becomes available during the “downhill” passage of electrons to oxygen:

NADH → NAD+  + H+  + 2e −  ∆G′ 0  = −158.2 kJ

(2.56)



1 O  + 2H+  + 2e −→H2O  ∆G′ 0  = −61.9 kJ 2 2

(2.57)

Overall reaction:

NADH + 

1 O2 + H+ →NAD+  + H2O  ∆G′ 0  ≈ −220 kJ 2

(2.58)

Refer to Equations 2.26 through 2.28 for FADH2 oxidation reactions. This free energy is conserved as electrochemical energy/proton motive force (pmf)/proton potential (∆p) as a result of proton translocation during electron transport. Electrochemical energy is a function of both the membrane potential (∆Ψ, electrical energy) and the difference in concentrations between solutions separated by the membrane (chemical energy). For n moles of protons that cross a membrane,

Electrochemical energy (∆μ) = Electrical energy + Chemical energy

(2.59)



Electrical energy = nF∆Ψ (J)

(2.60)

2-23

Catabolism and Metabolic Fueling Processes Glucose

2NAD+

ATP

2NADH

ADP

2

OAA 3

NADH

ADP

6

Malate

Acetyl-CoA

FADH2

Lactate

7

CoA

8

Fumarate

NAD+

NADH

ATP

Pyruvate

4

5

ATP PEP

Pi

NAD+

ADP

1

CO2

+

CO2

9

Formate

H2

Acetyl-CoA

FAD

Pi

Succinate

10

NADH

12

NAD+

CoA Acetyl-P

Acetaldehyde* NADH

ADP 11

13

NAD+

ATP

Acetate

CoA

14

NADH

Acetoacetyl-CoA Acetate 15

NAD+

22

β-OHButyryl-CoA

Ethanol

16 Crotonyl-CoA Pi

Butyryl-P ADP 21

ATP

CoA

17 20

NADH NAD+

Butyryl-CoA 18

NADH

Acetyl-CoA Acetoacetate 23

CO2

Acetone 24

NADH NAD+

Isopropanol

NAD+

Butyraldehyde NADH 19

Butyrate

NAD+ Butanol

Figure 2.7  Common fermentation pathways. Various fermentative products are produced from pyruvate in different organisms. Common products include succinate, lactate, formate, ethanol, acetate, acetone, butanol, isopropanol, and butyrate. Enzymes involved: 1, glycolytic enzymes; 2, PEP carboxylase; 3, malate dehydrogenase; 4, fumarase; 5, fumarate reductase; 6, pyruvate kinase; 7, lactate dehydrogenase; 8, pyruvate-formate lyase; 9, formate-hydrogen lyase; 10, phosphotransacetylase; 11, acetate kinase; 12, acetaldehyde dehydrogenase; 13, alcohol dehydrogenase; 14, acetyl-CoA-acetyl transferase; 15, L(+)-β-hydroxybutyryl-CoA dehydrogenase; 16, 1,3hydroxy-acyl-CoA hydrolase; 17, butyryl-CoA dehydrogenase; 18, butyraldehyde dehydrogenase; 19, butanol dehydrogenase; 20, phosphotransbutyrylase; 21, butyrate kinase; 22, CoA transferase; 23, acetoacetate decarboxylase; 24, isopropanol dehydrogenase. *Pyruvate can be nonoxidatively decarboxylated directly to acetaldehyde by the enzyme pyruvate decarboxylase.

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Cellular Metabolism

where F is Faraday’s constant and ∆Ψ is the potential difference over which the charge is moved. Also,

 [H+ ]  Chemical energy =  RTln  +in  (J)  [ Hout ] 

(2.61)

+ ] is the concentration of protons where [Hin+ ] is the concentration of protons inside the cell and [Hout outside the cell. The electrochemical energy or proton motive force is hence written as:



 [H+ ]  Δµ = nFΔΨ + RT ln  +in  (J)  [ Hout ] 

(2.62)

Electrochemical energy conserved in the proton gradient is expended when extruded protons are pumped backed into the matrix to power ATP synthesis through a membrane-spanning multisubunit protein called the F1F0 ATPase (ATP synthase). Electron transfer coupled to electrogenic translocation of protons through a membrane is referred to as vectorial translocation, and is achieved by electroncarrying proton pumps [6]. Examples of proton pumps include the cytochrome c oxidase complex in the mitochondrial ETC and the NDH-1 (NADH:ubiquinone oxidoreductase complex) in bacteria. In scalar translocation, protons are transferred from one side of the membrane to the other due to switches in electron carriers involved in the electron transport process [6]. For example, in the “Q cycle,” in which quinol (QH2) is oxidized to quinone (Q), electron carriers alternate between those that carry hydrogen and those that do not, such that protons are carried across the membrane and released when this switch occurs [6]. The “Q cycle” is illustrated in Figure 2.8. Complexes in the ETC where both electron transfer and proton translocation occur are called coupling sites. 2.6.5.1 Electron Transport in Mitochondria In eukaryotes, oxidative phosphorylation takes place in the mitochondria. The mitochondrial matrix contains high concentrations of electron transport enzymes, coenzymes, substrates and inorganic ions. In well-functioning mitochondria, oxidation of substrates and phosphorylation of ADP are tightly coupled and proceed simultaneously. The mitochondrial ETC is comprised of four complexes: NADH dehydrogenase/NADH:ubiquinone oxidoreductase (complex I), succinate dehydrogenase (complex II), ubiquinol:cytochrome c oxidoreductase (complex III), and cytochrome aa3 oxidase (complex IV). 2.6.5.1.1  Complex I NADH dehydrogenase is a large enzyme complex consisting of 42 different polypeptide chains and FMN and iron-sulfur proteins as its prosthetic groups [5]. This complex catalyzes the exergonic transfer of electrons from NADH to ubiquinone (“the Q pool”). Complex I is one of the three coupling sites in the mitochondria, translocating four protons (H+ ) across the membrane for every two electrons transferred [91].

NADH + Q + H+ →NAD+ + QH2    ∆E′ 0  = 360 mV ∆G′ 0  = −69.5 kJ/mole

(2.63)

2.6.5.1.2  Complex II FAD and iron-sulfur proteins are prosthetic groups in this complex and they funnel electrons derived from succinate oxidation into the Q pool via FADH2. No proton pumping occurs in this complex and free energy release is insufficient for ATP synthesis.

2-25

Catabolism and Metabolic Fueling Processes Intermembrane space

Inner membrane

QH2

Matrix QH2

2H+ + e– Q •– Q e– Hemes e– Q

Q

2H+

Q •– 2H+ + e– QH2 QH2

QH2

Q •– + e– + 2H+out

QH2 + Q •– + 2H+matrix QH2 + 2H+matrix

Q + QH2 + e– + 2H+out

Q + 2e– + 4H+out

Figure 2.8  Diagram of the mitochondrial Q cycle. In complex III, four protons are translocated across the inner mitochondrial membrane for every two electrons delivered from complex I in the form of a single reduced coenzyme Q (QH2). Two QH2 molecules diffuse to the outside of the membrane and each delivers a single electron to cytochrome c1 (via an iron-sulfur protein), forming two Q.− semiquinones and releasing a total of four protons to the intermembrane space. The two available Q.− electrons are delivered to hemes within the membrane, forming two oxidized coenzyme Q molecules which diffuse back to the matrix. Two matrix protons and two heme electrons are subsequently used to reduce one Q, regenerating one QH2.



Succinate + FAD→Fumarate + FADH2 FADH2 + Q → FAD + QH2    ∆E′ 0  = 85 mV ∆G′ 0  = −16.4 kJ/mole



(2.64)

(2.65)

2.6.5.1.3  Complex III Complex III is the second coupling site in the mitochondria. In this complex, oxidation of ubiquinol (QH2) by cytochrome c is coupled to the translocation of four protons. Electron carriers involved in the redox reactions include cytochrome b, cytochrome c1 and an iron-sulfur protein. Ubiquinol is a twoelectron carrier while cytochrome c is a one-electron carrier, and this switch is accommodated by the Q cycle (depicted in Figure 2.8). The overall reaction in complex III is:

 + QH2 + 2cyt c1 (oxidized) + 2H +matrix → Q + 2cyt c1(reduced) + 4H  intermembrane space

∆E′ 0  = 190 mV ∆G′ 0  = −36.7 kJ/mole

(2.66)

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Cellular Metabolism

2.6.5.1.4  Complex IV Complex IV transfers two electrons from cytochrome c to oxygen, while two protons are translocated across the membrane. Complex IV contains hemes a and a3 and three copper ions, which are used in electron transfer. Heme a3 is an oxygen-binding heme. The overall reaction in this complex is:  +    + 2cyt c(reduced) + 4H  matrix + ½O2 → 2cyt c(oxidized) + 2H intermembrane space + H 2O



∆E′ 0  = 580 mV ∆G′ 0  = −112 kJ/mole

(2.67)

Figure 2.9 is a schematic of the ETC in mitochondria. NADH oxidation in an intact mitochondrial ETC is accompanied by the transmembrane extrusion of a maximum of ten protons per mole of NADH or per oxygen atom reduced, while FADH2 oxidation is accompanied by the transmembrane extrusion of six protons per mole of FADH2 or per oxygen atom [6]. This ratio of protons extruded per oxygen atom reduced is referred to as H+/O. Intermembrane space

Innermembrane space

Matrix

Complex I ∆E´0= 360 mV ∆G´0 = –69.5 kJ/mole

NADH + H+ 2H FMN

NAD+

FeS-protein

4H+

4H+ Succinate FAD

2H

Fumarate

Coenzyme Q FeS

2H

Cyt bc1 4H + Complex III ∆E´0 = 190 mV ∆G´0 = –36.7 kJ/mole

2H +

Figure 2.9  Electron transport in mitochondria.

2H+

protein Cyt c

Complex II ∆E´0= 85 mV ∆G´0 = –16.4 kJ/mole

2e– 2H+

Cyt aa3 Complex IV ∆E´0 = 580 mV ∆G´0 = –112 kJ/mole

1 O + 2H+ 2 2 H 2o 2H+

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Catabolism and Metabolic Fueling Processes

The number of moles of ATP (cytosolic) synthesized per oxygen atom reduced (P/O ratio) in the mitochondria has been reported to range from ~0.96 to 2.9 for NADH oxidation and ~0.88−1.9 for succinate oxidation. There is evidence however that these ratios are close to 1.0, 1.0, and 0.5 for the three proton pumping sites [92]. Therefore the P/O ratio for NADH-linked substrates is 2.5 and 1.5 for succinate [91,92]. This is based on the consensus value of four protons for ATP synthesis: three protons driving the ATP synthase per ATP (3H + /ATP) and one for transport of Pi, ADP and ATP [5,91]. 2.6.5.2 Electron Transport in Bacteria The ETC in bacteria is found in the cytoplasmic membrane and is similar to, but more complex than, the mitochondrial ETC. The complexity of the bacterial ETC is attributed to the ability of bacteria to alter their electron transport mechanisms based on growth conditions or available electron donors and acceptors [6]. Bacterial ETCs can be divided into quinone-reducing branches and quinol-oxidizing branches. The reductive branches are responsible for electron transfer from electron donors to quinone and consist of the dehydrogenases, while the oxidative branches are redox systems involved in the oxidation of quinols, cytochromes and terminal oxidoreductases [93]. Figure 2.10 depicts a generalized ETC in bacteria. Depending on the organism, electrons entering the bacterial ETC can originate from a wide variety of donors including NADH, FADH2, organic substrates, H2, NH3, NO2− , sulfur, sulfide, and ferrous iron. The entrance point of electrons into the ETC depends on the reduction potential of the donor, with more negative potentials entering at higher “levels” of the ETC (NADH dehydrogenase being the highest level). Bacterial terminal electron acceptors include oxygen, nitrate, nitrite, fumarate, sulfate, carbon dioxide and dimelthylsulfoxide. Of these compounds, oxygen has the largest (positive) reduction potential and its reduction yields the largest Gibbs free energy. The well-studied ETC of E. coli is a good illustration of the complex and branched nature of bacterial ETCs. Proton translocation occurs during NADH oxidation (2H+/e −) via NADH:ubiquinone (a) Electron donor

Reductase(s) Dehydrogenase

(b)

bc1

Quinones

c

Terminal oxidase(s)

Electron acceptor

aa3

b

c

bc

aa3

Electron acceptor

Quinones

o

o b

o

Figure 2.10  Generalized bacterial electron transport chain. (a) Electrons from donors enter the ETC through dehydrogenase complexes, the quinone pool or the cytochromes. The ETC can branch at the quinones (quinols) and cytochromes. Under anaerobic conditions, electrons are transferred from the quinols to reductase complexes which reduce the final electron acceptor. Under aerobic conditions, electrons are transferred from quinols to terminal oxidases via multiple routes. (b) Examples of different electron transfer routes employed during aerobic respiration. Cytochromes o and aa3 are terminal oxidases.

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Cellular Metabolism

oxidoreductase (NDH-1), while a second NADH dehydrogenase (NDH-2) does not pump protons [6]. Several different quinol-oxidizing branches connect to terminal reductases or oxidases. Under aerobic conditions the cytochrome complexes bo and bd are formed for quinol oxidation [6]. Cytochrome bo has a lower affinity for oxygen compared to cytochrome bd, and their relative activities depend on the oxygen tension. Cytochrome bo translocates 2H +/e − (one scalar and one vectorial) while cytochrome bd translocates 1H +/e − (one scalar). During anaerobic growth, reductase complexes reduce electron acceptors. For example, nitrate reductase is synthesized for nitrate reduction and fumarate reductase for fumarate reduction. The complexity of the bacterial ETC makes it difficult to postulate a specific value for the P/O ratio, as it depends on the branches involved and the number of coupling sites. P/O ratios are higher in bacterial ETCs compared to mitochondria ETCs because protons are not required to bring Pi into the cell [6]. 2.6.5.3 Efficiency of Energy Conversion The efficiency of a process is defined as the ratio of energy output to energy input. The efficiency of electron transport in converting free energy to ATP is given as: ξ=



N × E ATP ×100 E react

(2.68)

where ξ is the efficiency, N is the number of moles of ATP produced, EATP is the energy in one highenergy phosphoanhydride bond in ATP or the free energy from ATP hydrolysis, and Ereact is the energy released from the transport process.

Example 1:  Efficiency of Oxidative Phosphorylation Oxidation of one mole of NADH in mitochondria under standard biochemical conditions (298 K, 1 atm, pH 7.0), has a theoretical yield of 220 kJ of free energy per mole of NADH. If the mitochondrial P/O ratio is taken as ~2.5, the efficiency of oxidative phosphorylation in mitochondria under standard biochemical conditions is calculated as:



NADH + 

1 O  + H+ →NAD+  + H2O   ∆G´0 ≈ −220 kJ/mole 2 2

ADP + Pi ↔ ATP + H2O    ∆G´0 = +30.5 kJ/mole ξ = -2.5 ×

30.5 × 100  ≈ 35% 220

(2.69) (2.70) (2.71)

Example 2:  Efficiency of Glucose Catabolism Under standard biochemical conditions, complete oxidation of glucose to CO2 has a theoretical maximum yield of 2,840 kJ of energy per mole of glucose [5]. If 32 moles of ATP are produced by substrate level phosphorylation and oxidative phosphorylation, then the efficiency of glucose oxidation is:



ξ = -32 ×

30.5 × 100  ≈ 34% 2840

(2.72)

Actual values for efficiency differ from these calculated values since cells do not operate under standard biochemical conditions.

Catabolism and Metabolic Fueling Processes

2-29

2.6.6  Key Parameters Influencing Regulation of Catabolic Pathways Metabolic activities are regulated to ensure a balance between catabolism and anabolic demands. Catabolic pathways are sensitive to and tightly regulated in response to a number of related signals that reflect the redox potential and energy charge of the cell. This includes the presence of oxygen or other terminal electron acceptors and whether a growth substrate is limiting or in excess. As was described in Section 2.6.2 for the regulation of PDHc and PFL enzymes (involved in pyruvate metabolism), key “control” enzymes of the central pathways are allosterically regulated by central pathway intermediates. While the effectors vary between isozymes and organisms, some general examples include: inhibition of phosphofructokinase by ATP and PEP, activation of pyruvate kinase by fructose-1,6-bisphosphate, inhibition of citrate synthase and isocitrate dehydrogenase by ATP and NADH, and inhibition of α-ketoglutarate dehydrogenase by its products NADH and succinyl-CoA. Transcription factors responsive to the intracellular redox potential also play an important role in regulating catabolic pathway genes. The most characterized are the fumarate nitrate reductase (FNR) and the ArcA/B global regulatory systems in E. coli (reviewed in Ref. [6]). FNR is minimally functional during aerobic growth, but under anaerobic or very low oxidation-reduction potential conditions, FNR acts as a positive regulator of many genes including PFL and fumarate reductase. FNR additionally represses many other aerobically expressed genes such as succinate dehydrogenase (in the TCA cycle) and respiratory cytochromes. Transcriptional control exerted by the Arc system also includes induction as well as repression and has pronounced effects on central metabolism under microaerobic conditions (where FNR may still be relatively inactive) in addition to anaerobic conditions, in coordination with FNR (many genes are regulated by both FNR and Arc) [94–96]. As examples, the Arc system is responsible (at least partly) for repression of PDHc and TCA cycle genes and induction of PFL. “Catabolite repression” in microbes refers to attenuated activity of inducible enzymes involved in the catabolism of a specific carbon source (i.e., the inducer, such as xylose or arabinose) when there is a surplus of a more “favorable” substrate such as glucose [97,98]. Cyclic AMP (cAMP) acts as an effector of the “catabolite activator protein” (CAP) transcription factor (also called cAMP-receptor protein, or CRP), which regulates transcription of many (or most) catabolic operons [97,99–102]. Glucose metabolism lowers intracellular cAMP concentrations [103] and as a result glucose is preferentially metabolized when present with other substrates whose catabolic genes are under CAP control. Microorganisms thus often exhibit “diauxie” or “diauxic growth” in the presence of sugar mixtures, a phenomenon in which multiple carbon sources are not metabolized simultaneously but rather utilized sequentially [98,104– 106]. Elimination of diauxic growth has been achieved in engineered E. coli strains by elevating the cAMP pool [105,107] and expressing a constitutive CRP mutant [108]. In the presence of excess nutrients, organisms often exhibit “overflow metabolism” in which the carbon substrate is assimilated rapidly but incompletely oxidized, resulting in production of some ATP and biomass as well as secretion of overflow metabolites such as ethanol and acetate (even in the presence of ample oxygen) [109–112] Respiration essentially does not “keep-up” with glycolytic flux and glycolysis is not attenuated, so metabolites are secreted to maintain redox balance and ATP production (much like fermentation) [35,111,113–116]. When the high-energy nutrients have been exhausted, cells switch to a survival agenda in which they assimilate the previously excreted organic compounds. Examples of organisms which exhibit overflow metabolism in the presence of excess glucose include E. coli (acetate accumulates and is later metabolized in what is referred to as the “acetate switch”) [117], Saccharomyces cerevisiae (ethanol accumulates) [111,118,119], and Klebsiella aerogenes (excretes 1,3-propanediol) [120]. As noted, the TCA cycle is subject to multi-tiered regulation at the levels of transcription and enzyme activity [64,65,121,122]. Under anaerobic conditions enzyme activities are 10- to 20-fold lower than those during aerobic growth [51,64,66,121,123]. High catabolic flux caused by excess glucose results in repression of TCA cycle/respiratory pathways (the “Crabtree” effect), influenced by a combination of factors including reduced gene expression (catabolite repression) and inhibition by elevated levels of “energetic” metabolites [64,113,122,124]. From a metabolic engineering perspective, it is often desirable

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Cellular Metabolism

to alleviate this repression and increase flux through the TCA cycle or specific TCA cycle enzymes. This has been demonstrated through deletion of the redox-sensitive regulators (ArcA and FNR) [61,113] and through use of a constitutive CRP mutant [108].

2.7  Concluding Remarks While not comprehensive, this overview of catabolism and metabolic fueling processes describes the pathways of primary concern to metabolic engineers. Although innumerable variations are found throughout nature, the basic energy conservation mechanisms described here are used by all organisms. For example, while we have not described the roles of nonproton ion gradients in cellular energetics/ATP synthesis (a crucial component of methanogen metabolism), or a variety of unique features of archaeal respiratory chains [125], very similar biomolecular components and energy coupling mechanisms exist, and the same thermodynamic relationships apply. An understanding of these fundamental, naturally devised processes that harness energy for life should help to facilitate approaches to engineer strains with improved properties (e.g., to overcome growth deficits in engineered strains or redirect high-energy intermediates toward energy-dependent pathways of interest). Knowledge of the metabolic couplings between growth, ATP production and redox balancing is additionally necessary for constraining metabolic networks through genetic engineering, to develop genetic selections that guide the design of strains with unique phenotypes. An ever-growing need for sustainability through renewable and biologically derived products brings metabolic engineering to the technology forefront. In this context, bioprocess efficiency is of central importance; with regard to distribution of carbon and energy, it is not difficult to envision scenarios in which an optimal tradeoff exists between cell growth and product formation. Vital to our success in this arena is an ability to restructure catabolic pathways, made possible by the genomic revolution and the limitless insights and tools that it provides.

Acknowledgment The authors thank the National Science Foundation for financial support (grant No. BES0519516).

References 1. Woese, C.R., O. Kandler, and M.L. Wheelis. Towards a natural system of organisms—proposal for the domains archaea, bacteria, and eucarya. PNAS, 87 (12), 4576–4579, 1990. 2. Purves, W.K., et al. Life: The Science of Biology. 6th ed. Sunderland, MA: Sinauer Associates Inc., 2001. 3. Todar, K. The diversity of metabolism in prokaryotes. In Todar’s Online Textbook of Bacteriology, 2004. www.textbookofbacteriology.net/metabolism.html 4. Ingraham, J.L., O. Maaloe, and F.C. Neidhardt. Growth of the Bacterial Cell. Sunderland, MA: Sinaeur Associates Inc., 1983. 5. Nelson, D. and M. Cox. Lehninger Principles of Biochemistry. 4th ed. New York: W.H. Freeman and Company, 2005. 6. White, D. The Physiology and Biochemistry of Prokaryotes. 2nd ed. New York: Oxford University Press, 2000. 7. Jackson, J.B. The proton-translocating nicotinamide adenine dinucleotide transhydrogenase. J. Bioenerg. Biomembr., 23 (5), 715–741, 1991. 8. Bizouarn, T., et al. The involvement of NADP(H) binding and release in energy transduction by proton-translocating nicotinamide nucleotide transhydrogenase from Escherichia-coli. Biochim. Biophys. Acta Bioenergetics, 1229 (1), 49–58, 1995. 9. Hutton, M., et al. Kinetic resolution of the reaction catalyzed by proton-translocating transhydrogenase from Escherichia coli as revealed by experiments with analogs of the nucleotide substrates. Eur. J. Biochem., 219 (3), 1041–1051, 1994.

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3 Biosynthesis of Cellular Building Blocks: The Prerequisites of Life 3.1

Introduction ���������������������������������������������������������������������������������������3-1

3.2

Amino Acid Biosynthesis �����������������������������������������������������������������3-3

3.3

Nucleotides as Building Blocks �����������������������������������������������������3-12

3.4

Synthesis of Carbohydrates for Building Cells........................... 3-20

3.5

Cell Synthesis of Lipids ������������������������������������������������������������������ 3-25

Incorporation of Biological Atoms into the Cells  •  Cellular Building Blocks Are Derived from Central Precursors The 2-Oxoglutarate Derived Amino Acids  •  The Aspartate Family and Branch-Chained Amino Acids  •  Serine-Glycine Family of Amino Acids  •  Aromatic Nonpolar Amino Acids  •  Histidine Biosynthesis Structure and Organization of Nucleotides  •  Biosynthesis of Nucleotides  •  Nucleotide Metabolic Conditions

Zachary L. Fowler, Effendi Leonard, and Mattheos Koffas State University of New York at Buffalo

Introduction to Carbohydrates  •  Interconversion of Carbohydrates  •  Other Import Enzymes of Carbohydrate Synthesis Introduction to Lipids and Fatty Acids  •  Sources of Metabolites for Lipogenesis  •  Synthesis of Long-Chain Fatty Acids  •  Synthesis of Neutral Lipids

References ����������������������������������������������������������������������������������������������������3-31

3.1  Introduction The vast array of secondary metabolites critical for the production of commercial pharmaceuticals, food products, and other consumer goods are all synthesized from the most basic cellular building blocks. This chapter covers the fundamentals of their biosynthesis, where a majority of the biosynthetic routes have been obtained from studies of well characterized prokaryotes, especially the enteric bacterium Escherichia coli. However, especially in the cases of amino acid and nucleotide biosynthesis, it is safe to assume all organisms have a universal cellular biochemistry thus generalizing the fundamental (primary) biosynthetic pathways as similar in all organisms to some extent. Amino acids, nucleotides, sugars, and fatty acids make of the major building blocks used in the synthesis of macromolecular cellular components including protein polymers, nucleic acids (deoxyribonucleic acid, DNA, and ribonucleic acid, RNA), cellular structures, and lipids. Cellular building blocks are typically assembled out of degradation products from central precursor metabolites, specifically acetyl-CoA, pyruvate, oxaloacetate, ­2-oxoglutarate, phosphoenolpyruvate (PEP), and the hexose phosphates, which themselves are derived from the organic, simple inorganic, low-molecular weight, and macromolecular molecules. Cells are constantly shifting from metabolic states to catabolic states, generating and consuming the building blocks 3-1

3-2

Cellular Metabolism

in an effort to maintain the delicate equilibrium essential to proper cellular functioning while growing on environmental substrates. Because most growth substrates eventually lose nitrogen, phosphorus, and sulfur during degradation processes, the central precursor metabolites contain only carbon, hydrogen, oxygen, and some phosphate atoms. As a result, cells also require compounds rich in nitrogen, phosphate, sulfate, one carbon units, as well as energy (mostly in the form of phosphoric acid anhydride bonds in adenosine triphosphate (ATP)) and a reductant in the form of the hydride of NADPH.

3.1.1  Incorporation of Biological Atoms into the Cells 3.1.1.1 Nitrogen Nitrogen is incorporated into a cell’s metabolism in several oxidation states including NH+ 4 , NO3− , N2, as well as through urea and amino acids, and purine and pyrimidine metabolites, however the most common route for utilization of inorganic nitrogen is generally ammonia. Reactions involving nitrogen assimilation in all organisms lead to the formation of central nitrogen carriers, primarily glutamate, and glutamine, and to a lesser extent the secondary metabolites aspartate and carbamoyl phosphate. Carbamoyl phosphate is unique in that it is only utilized for the biosynthesis of arginine, urea, and pyrimidine nucleotides. As such most nitrogen transfer from ammonia to the amino acids and other nitrogen containing molecules is conducted through the two primary nitrogen carriers’ glutamate and glutamine. By and large, the process from substrate uptake to cellular component synthesis using nitrogen is carried out in three steps. First, and only after nitrogen sources from the environment are transported into the cells, ammonia formation occurs through the reduction of nitrite or N2 or by hydrolytic release mechanisms from nitrogen containing organic substrates. The second step involves nitrogen transfer, in which case, the ammonia is transferred to form glutamate and glutamine. Glutamine is the direct nitrogen donor for most nitrogen containing building blocks, except for the α-amino group of the amino acids for which glutamine is only an indirect donor, where glutamate on the other hand, is the universal amino donor of α-amino acids. In E. coli ammonia assimilation is conducted through glutamine synthesis mediated by L-glutamine synthetase when not under energy limitation or by a glutamate dehydrogenase (GDH) pathway when energy in the cell is at a premium and phosphate levels are high. The GDH pathway is the primary ammonia assimilation route in fungi and yeast. In the third step, nitrogen is finally released by transferring the α-amino group of L-glutamate to 2-oxo acids, the direct precursors of α-amino acids, by aminotransferase enzymes. In general, most cell types possess a variety of aminotransferases in which each enzyme is uniquely responsible for the synthesis of a group of 2-oxo acids. 3.1.1.2  Sulfur Elemental sulfur, which can be synthesized from microbial or air oxidation has a low solubility; therefore, it is only used by a few prokaryotes as an electron acceptor in aerobic respiration. On the other 2− hand, sulfate (SO2− 4 ) and thiosulfate (S2O3 ) are the principal sulfur sources for prokaryotes. Under aerobic respiration sulfur sources are commonly transported into the cells using a process mediated by an ATP-dependent, high-affinity, binding-protein-dependent (ABC) transport system while the low affinity transport systems is typically inactive even if present. Anaerobic respiration results in sulfate reduction to H2S by a variety of sulfate reducing enzymes, a process that has become highly evolved in geothermal bacteria. In many cases cellular metabolism finds itself in a condition between aerobic and anaerobic where thiosulfate and H2S (the preferred substrates of anaerobic bacteria) are both present. H2S is highly toxic to most cells; therefore it is immediately transferred into the amino acid cysteine by the enzyme O-acetylserine sulfhydrylase. Intracellular sulfur containing compounds, for instance ­methionine, coenzyme A, lipoic acid, thiamine pyrophosphate, and glutathione, ultimately obtain their sulfur from cysteine. Through a similar enzyme, thiosulfate is also incorporated into cysteine forming cysteine thiosulfate. While the transport of sulfate and synthesis of cysteine are in no way linked, both are controlled by the complex cys regulon in that the presence of excess cysteine results in feedback-inhibition of O-acetylserine formation, which is a critical inducer of expression for the sulfate reducing enzymes.

Biosynthesis of Cellular Building Blocks: The Prerequisites of Life

3-3

3.1.1.3  Phosphate Since it is readily oxidized by air, phosphorus exists mostly in the form of phosphate, which is the main inorganic phosphorus source of prokaryotes and does not need further reduction. The two phosphate forms that exist in neutral pH environments, H2PO−4 and HPO2− 4 are readily transported into the cells by unique transport systems. At high concentrations (>0.1 mM), a constitutively expressed, relatively unspecific, low-affinity system driven by proton motive force through proton symport operates with high capacity to alleviate the cell from excess phosphorus levels. When phosphate exists in low ­concentrations (<10 µM), transport into the cells is governed by a well regulated, high-affinity binding-protein-dependent system driven by the hydrolysis of ATP. Once transported into the cells, phosphate is transferred to ADP by ATP synthase to form ATP, the universal phosphate carrier and cellular energy source. 3.1.1.4 One-Carbon Units and Oxygen The term one-carbon (C1) unit refers to carbon atoms separated from other carbon atoms by hetero atoms such as oxygen, nitrogen, and sulfur. Biosynthesis of C1 units originates from coenzymes containing C1 units bound to a nitrogen or sulfur atom. In most cases, C1 units are synthesized from serine or glycine through serine hydromethylase or glycine decarboxylase, respectively to form the conjugation with tetrahydrofolate (H4F) forming methylene-H4F. Methylene-H4F is either reduced to methyl-H4F or successively oxidized to the formyl level where these C1-H4F compounds serve as C1 precursors in a number of reactions, for example purine and methionine biosynthesis. In general, molecular oxygen is not required in biosynthetic reactions because all cellular oxygen essentially, is obtained from water. However, O2 is required in a few hydroxylation reactions or in the introduction of double bonds performed by some obligate bacteria to synthesize uniquinone of monosaturated fatty acids and polysaturated fatty acids.

3.1.2  Cellular Building Blocks Are Derived from Central Precursors The three major metabolic pathways serving as suppliers of central precursor metabolites for the synthesis of building blocks are glycolysis, the citric acid cycle and the pentose phosphate pathway, each of which has multiple, tightly regulated steps that will not be discussed here. Glycolysis and the reverse process gluconeogenesis are used to generate the C6 carbon base molecules for many of the secondary metabolites with functional phosphates attached. The citric acid cycle (also known as the tricarbocyclic acid cycle (TCA)) is the energy pump for aerobic respiration generating ATP to such amounts that the complete oxidation of pyruvate to CO2 and H2O by the TCA cycle produces 18 ATP equivalents. Also known as the Calvin cycle in plants, the pentose phosphate pathway is used to convert the C6 carbons into C5 carbons, ribose 5-phosphate and the other C3, C4, C5, and C7 sugars. In some bacteria, alternative metabolic pathways exist to provide the essential central precursor metabolites, for example, the Entner–Doudoroff pathway. The synthesis of cellular building blocks through the degradation of central precursor metabolites is strictly regulated and involves anaplerotic reactions to replenish the metabolic constituents.

3.2  Amino Acid Biosynthesis All enzymatic metabolic function is conducted by proteins making them the major cell constituents synthesized within the cell. As such the generation of the 20 natural L-amino acid building blocks using only a few central precursor metabolites for their biosynthesis to form proteins is a critical process (Figure 3.1). Additionally, L-amino acids serve as precursors for the synthesis of nucleotides. Nitrogen containing metabolites can be formed directly from a single nitrogen source, such as ammonia or nitrate in many bacteria and most plants; however preformed amino acids are often the substrates of other nitrogen containing metabolites in many organisms. Mammals synthesize only half of the 20 amino

3-4

Cellular Metabolism Pentose phosphate Proteins

Glycolysis / Gluconeogenesis Glucose 6-phosphate

Ribose 5-phosphate

Fructose 6-phosphate

Amino acids Erythose 4-phosphate

Glycerol Glyceraldehyde 3-phosphate

Nucleic acids Lipids 3-Phosphoglycerate Nucleotides Phosphoenolpyruvate

Fatty acids

Mono/polysaccharides Pyruvate

Acetyl-CoA Citrate cycle (TCA cycle)

Oxalacetate 2-Oxoglutarate Succinate

Figure 3.1  Interconnections in the formation of building blocks. The figure highlights the interconnectivity between metabolic pathways producing the critical cellular constituents needed to maintain cell growth. Only those intermediates critical to the synthesis of building blocks are noted in glycolysis, the TCA cycle, and the pentose phosphate pathway.

acids required for growth and maintenance; these amino acids are termed “non essential amino acids” with the remaining “essential amino acids.” Generally, nonessential amino acids are readily synthesized from metabolites derived from the central metabolism, such as glycolysis or the citric acid cycle. On the other hand, essential amino acids which generally exhibit complex structures such as aromatic rings and hydrocarbon side chains need to be provided from diets. Amino acids can be categorized into five biosynthetic families plus histidine, based on the central metabolic precursors.

3.2.1 The 2-Oxoglutarate Derived Amino Acids Degradation of the metabolite 2-oxoglutarate leads to the synthesis of glutamate, glutamine, proline, and arginine in which two primary reactions, amidation, and reduction, are utilized for the synthesis of this amino acid class (Figure 3.2). As mentioned before, glutamate is the universal

ATP

NH3

C

C

CH2

CH2

C

NH2

H

COOH

1

1

Glutamine

O

H2N

Pi

C H2

H Glutamate

C H2

ADP

C

NH2

ATP

COOH

ADP

2

10

–

11

2 [H]

Pro

Arg

3

Pi

4

ATP ADP 2 [H]

H

O C

Pi

C H2

C

2 [H]

H2O

NH

C

CH3

C H2

C

NH2

Proline

HN

COOH

12

COOH

5

Glutamate

COOH C C H2 H Semialdehyde

C H2

O

H Semialdehyde

H

O

Spontaneous

6

C

ATP

HN

C

HN

C H2

C H2

NH2

H2 C

Fumarate

C H2

NH2

H

C

NH2

COOH

COOH

NH2

Arginine

C C H2 H

9

8

COOH

Aspartate

Citrulline

C C H2 H

7

Ornithine

C H2

NH2

H2 C

Pi

CAP

H2 C

AMP + PPi

O

HN

H2N

H2O Acetate

Figure 3.2  Glutamate family amino acids biosynthetic pathways. Glutamate (Glu) is formed by reductive amination of  2-oxoglutarate. Glutamine (Gln) is formed by then using: 1, Glutamate synthase. Synthesis of arginine (Arg) requires: 2, N-acetylglutamate synthase; 3, N-acetylglutamate kinase; 4, N-acetylglutamyl-phosphate reductase; 5, N-acetylornithine-δ-transaminase; 6, N-acetylornithinase; 7, ornithine carbamoyl-transferase (or isozymes); 8, argininosuccinate synthetase; 9, argininosuccinase. For the path to Proline (Pro): 10, γ-glutamyl kinase; 11, glutamate-γ-semialdehyde dehydrogenase; 12, Δ1-pyrroline-5-carboxylate reductase.

HO

O

Acetyl -CoA

CoA -SH

–

2-Oxoglutarate

Biosynthesis of Cellular Building Blocks: The Prerequisites of Life 3-5

3-6

Cellular Metabolism

nitrogen donor for α-amino groups in amino acids. The synthesis from 2-oxoglutarate through reversible reductive amination is catalyzed by the enzyme GDH, which is the primary route in bacteria utilizing ammonia as the sole carbon source. However, in animal cells, the primary role of GDH (located in mitochondria) is for the synthesis of 2-oxoglutarate in the citric acid cycle for energy generation. Moreover, the enzyme is allosterically regulated, in which 2-oxoglutarate synthesis is activated by ADP and GDP, and is inhibited by ATP and GTP. In this case, glutamate synthesis from 2-oxoglutarate and glutamine is catalyzed by glutamate synthase. In general, glutamate synthase serves as the primary enzyme for glutamate synthesis in most cells since GDH has a relatively low affinity for ammonia. Glutamine is synthesized by the attachment of a second ammonia moiety to glutamate by the enzyme glutamine synthetase, which requires ATP-dependent glutamyl-phosphate formation in order to eliminate the −OH group and to drive the amidation reaction forward. Glutamine is a universal nitrogen carrier with multiple key roles, in which its amide nitrogen is used for the synthesis of glutamate, tryptophan, histidine, nucleotides, and amino sugars. For this reason, its biosynthesis is under tight regulation. In E. coli, glutamine synthetase is regulated by two unique interdependent mechanisms. One regulatory mechanism controls the allostericity of the enzyme by cumulative feedback inhibition in which the allosteric regulation is accommodated by the presence of eight binding sites for eight inhibitor metabolites derived from glutamine metabolism, i.e., glucosamine-6-phosphate, carbamoyl phosphate, CTP, AMP, and the amino acids alanine, tryptophan, histidine, and glycine. The inhibitory mode is dependent on the number of metabolites bound to the recognition sites such that a single inhibitor is not sufficient enough to halve glutamine biosynthesis. The other mechanism controls enzyme activities through covalent modifications by adenylation. There are 12 adenylation sites on glutamine synthetase which are located adjacent to the catalytic sites. At each site inactivation is achieved when specific tyrosine residues react with ATP to form esters at each site where, in a manner similar to the allosteric control, partial adenylation results in partial inactivation of the protein. The processes of adenylation and deadenylation are further mediated by a complex regulon with both reactions catalyzed by the formation of an enzyme complex of adenylyl transferase and a regulatory protein PII. The cascade provides a feedback loop that when glutamine is present in high concentration, the biosynthesis is shut down. On the other hand glutamine synthetase is turned on in the presence of low glutamine concentration by the accumulation of 2-oxoglutarate and with the presence of ATP. The biosynthesis of proline from glutamate is mediated by several steps. In the first step, glutamate is converted into N-acetylglutamate by the enzyme γ-glutamyl kinase. Next, N-acetylglutamate is converted into glutamic γ-semialdehyde by the enzyme glutamate γ-semialdehyde. Subsequently, glutamic γ-semialdehyde is spontaneously converted into 1-pyrroline carboxylic acid through an intramolecular Schiff base reaction, and further NADPH reduction results in proline. Proline synthesis is regulated by a negative feed-back inhibition of γ-glutamyl kinase. Similar to proline, arginine is synthesized from glutamate through a series of biochemical reactions, involving the enzymes N-acetylglutamate kinase, N-acetylglutamyl-phosphate reductase, N-acetylornithine-δ-transaminase, N-acetylornithase, ornithine carbamoyl-transferase, arginosuccinate synthetase, and argininosuccinase. Like proline, to modulate its synthesis from glutamate, arginine exerts a negative feedback inhibition to the enzyme N-acetylglutamate kinase.

3.2.2 The Aspartate Family and Branch-Chained Amino Acids Oxaloacetate serves as a central metabolic precursor for the synthesis of several amino acids making up the aspartate family. With glutamate, oxaloacetate undergoes a transamination reaction to form the central amino acid of aspartate (Figure 3.3). Subsequently, the conversion of aspartate to form asparagine is catalyzed by L-asparagine synthetase, which exhibits similar mode of action to that of glutamine synthetase. Initiating the pathway for the synthesis of lysine, threonine, methionine, and isoleucine, aspartate kinase I, II, III first converts aspartate to β-aspartyl phosphate. Next, β-aspartyl phosphate

3-7

Biosynthesis of Cellular Building Blocks: The Prerequisites of Life

COOH

Glutamate

C

H2N

Oxaloacetate

ATP

H

CH2

1a,b,c

ADP

COOH H2N

CH2

– – –

C O OH Aspartate

Thr Met Lys

ATP 10a,b

C O OP Aspartyl-4-P 2 [H] 2

PPi

H3C

Pi

NH3 or Glutamine

COOH

10a,b H2N

AMP

C

C

H 11

O H Aspartate semialdehyde

CH2 C

2 [H]

O NH2 Asparagine

O

2 H2O

C

H

C

COOH Pyruvate

CH2

COOH H2N

H

C

N

HOOC

COOH

Dihydrodipicolinate

3a,b

2 [H] 12

COOH H2N

Succinate Cystine H2N

C H CH2 S

H C

C

CoA -SH

H

CH2

5

Cys

COOH

H

H Cystathionine

O

C

Succinyl

H O-Succinyl homoserine

H2O

COOH

Succinyl -CoA H N 2 4

C

–

C

Met

CH2

H

ATP 8

HOOC N COOH Tetrahydrodipicolinate Succinyl -CoA 13 CoA -SH Glutamate 14

ADP Pyruvate + NH3 COOH C

H

CH2 H

OH

OH Homoserine

6

H2N

H

C

SH

H Homocysteine

H2O

COOH CH3-H4F 7

9

H2N C H CH2 H

C

S

Pi COOH CH3

H Methionine To branched-chain amino acids

H2N

C

H

H

C

OH

H

C

H

H Threonine

H2O

15

Succinate

16a,b H2C H2C H

CO2 NH2

CH2 C

NH2

COOH Lysine

Figure 3.3  Biosynthesis of the aspartate family of amino acids. Transamination of oxaloacetate leads to aspartate (Asp). The common pathway to threonine (Thr), methionine (Met), and lysine (Lys) includes: 1a/b/c, aspartate kinases I, II, and III; 2, aspartate semialdehyde dehydrogenase; 3a/b, homoserine dehydrogenase I, II (bifunctional enzymes). Methionine branch: 4, homoserine succinyltransferase; 5, cystathionine γ-synthase; 6, cystathionine β-lyase; 7, homocysteine methylase. Threonine branch: 8, homserine kinase; 9, threonine synthase. Asparagine (Asn) synthesis uses 10a/b, asparagine synthetases. Lysine branch: 11, dihydrodipicolinate synthase; 12, dihydrodipicolinate reductase; 13, tetrahydrodipicolinate succinylase; 14, succinyl diaminopimelate; 15, succinyl diaminopimelate desuccinylase; 16a/b, diaminopimelate epimerase/decarboxylase.

3-8

Cellular Metabolism

is transformed into aspartate semialdehyde by the enzyme aspartate semialdehyde synthase. Through homoserine dehydrogenase I, II (bifunctional enzyme aspartate kinases I, II), homoserine is next ­synthesized from aspartate semialdehyde. Further reduction occurs to alcohol homoserine by homoserine kinase followed by isomerization by threonine synthase to result in threonine. The biosynthesis of threonine is regulated through the inhibition of aspartate kinase I by threonine itself. Lysine is formed from the condensation (Schiff’s base formation) of aspartate semialdehyde and another central precursor metabolite, pyruvate, by the enzyme dihydrodipicolinate synthase to form dihydrodipicolinate. Further reduction catalyzed by dihydropicolinate reductase gives tetrahydrodipicolinate. Subsequent reactions catalyzed by tetrahydrodipicolinate succinylase, succinyl diaminopimelate aminotrasnferase, succinyl diaminopimelate desuccinylase, diaminopimelate epimerase, and diaminopimelate decarboxylase lead to the synthesis of lysine. Lysine also serves a negative inhibitor by acting on aspartate kinase III. Methionine biosynthesis branches from homoserine through the action of homoserine succinyltransferase to synthesize O-succinyl homoserine. Cystathionine formation then follows by a reaction catalyzed by cystathionine γ-synthase, a pyridoxal-phosphate enzyme, and also results in the release of succinate, an intermediate in the TCA cycle. Subsequent reactions involve cystathionine β-lyase that synthesizes homocystein and then homocystein methylase that catalyzes the incorporation of methylH4F, a C1 carrier, to the homocystein molecule generate methionine. Methionine is a feedback inhibitor of two enzymes; its own biosynthetic pathway at the step catalyzed by homoserine succinyltransferase and also aspartate kinase II found earlier in the aspartate family biosynthesis. While alanine is derived from the direct transamination of pyruvate, the biosynthesis of branchedchain amino acids isoleucine, valine, and leucine is more complicated but with many common principles between each path (Figure 3.4). Activated acetaldehyde or 2-oxobutyrate (derived from threonine deamination) is condensed with pyruvate by acetohydroxyacid synthases in a thiamine-pyrophosphatedependent reaction. The α-acetyl products are then reduced to 2,3-dihydroxyl compounds through catalysis by acetohydroxyacid isomeroreductase, followed by the elimination of a water molecule by dihydroxyacid dehydratase to yield 2-oxo compounds. Further transamination results in formation of valine and isoleucine. The formation of α-isopropylmalate from acetyl-CoA and 2-oxoisovalerate, an intermediate of the valine pathway, occurs by isopropylmalate synthase; this reaction initiates the ­biosynthesis of leucine. Subsequent reactions form 2-oxoisocaproate by isopropylmalate isomerase and β-isopropylmalate dehydrodenase, which is then followed by a transamination reaction resulting in the formation of leucine. The biosynthesis of leucine, valine and isoleucine is regulated by negative feedback inhibition to acetohydroxyacid synthases by valine and isoleucine.

3.2.3  Serine-Glycine Family of Amino Acids The biosynthesis of the serine-glycine family (Figure 3.5) from 3-phosphoglycerate originates from the oxidation of the free hydroxyl group by the enzyme 3-phosphoglycerate dehydrogenase. Subsequently, transamination and phosphate removal by 3-phosphoserine aminotransferase and 3-phosphoserine phosphatase result in serine biosynthesis. From here the biosynthetic pathway branches in two directions, one to cysteine by incorporation of hydrogen sulfide (HS) and the other into glycine. Biosynthesis of cysteine occurs through the action of the two enzymes serine transacetylase and O-acetylserine sulfhydrylase. In general, serine transacetylase catalyzes the condensation of serine with acetyl-CoA to give β-O-acetylserine. Which when HS is directly incorporated into by the enzyme O-acetylserine sulfhydrylase, results in cysteine. Moreover, the biosynthesis of cysteine is also regulated by the cys regulon. Mammals derive cysteine from diet or the metabolism of methionine because under normal physiological conditions, cysteine does not exist in cells as a free amino acid due to quick reduction by glutathione. Here, the route of cysteine biosynthesis proceeds through the reverse pathway of methionine. More specifically, serine and homocysteine are condensed by the enzyme cystathionine synthase to form cystathionine from which cysteine is then formed through the cleavage of cystathionine. On the

3-9

Biosynthesis of Cellular Building Blocks: The Prerequisites of Life O 1

CH2

NH3

H

O COOH

C

CH3

Pyruvate x2 2a,b,c

2a,b,c CO2 H3C

O

OH

C

C

COOH

CH2

H3C

CH2 H CH3 α,β-Dihydroxyβ-methylvalerate

H3C

Glutamate CH3

H3C

C

H

CH2

COOH

C

O

COOH 2-Oxoisocaproate 2 [H] CO2

2 [H] 8 7

C

C

CH3 COOH

CH3 H

H3C

C

H

HOOC

C

OH

α,β-Dihydroxyisovalerate

H2O

CH2 COOH α-Isopropylmalate

H2O CoA -SH

O

H C

C

COOH

CH2 α-Oxo-β-methylvalerate Glutamate 5 a,b H

NH2

C

C

CH2 H CH3 Isoleucine

H3C

H

O

C

C

COOH

H2O COS

2-Oxoisovalerate

H3C CoA Acetyl-CoA

Glutamate

5 a,b

H3C

6

COOH

CH3

CH3

H3C

C

4

4

H3C

C

5 c,d

OH OH COOH

C

OH

3

2 [H]

OH OH C

O

α-Acetolactate

α-Acetoα-hydroxybutyrate

H3C

Val Ile

CH3

CH3

3

COOH

C C H2 H

CH3 Leucine

CH3

2-Oxobutyrate

NH2

C

H3C

COOH

C

– –

Threonine

H

NH2

C

C

COOH

CH3 H Valine

Figure 3.4  Biosynthesis of the branched-chain amino acids. Parallel path for synthesis of isoleucine (Ile) and valine (Val): 1, threonine deaminase; 2a/b/c, acetohydroxyacid synthases (a/b are end-product inhibited); 3, acetohydroxyacid isomeroreducaste; 4, dihydroxyacid dehydrase; 5a/b, transaminases; 5c/d, transaminases; 6, isopropylmalate synthase; 7, isopropylmate isomerase; 8, β-isopropylmalate dehydrogenase.

3-10

Cellular Metabolism COOH C

O

CH2O P 2 [H]

Hyroxypyruvate 3-phosphate

COOH H

C

Glutamate

2

1 OH

CH2O P

Pi

COOH

Glycerate 3-phosphate

H2N 6

C

CH2OH H4F

COOH C H

H

Glycine 7

H2N

C

H

CH2O P Serine 3-phosphate

3 Acetyl -CoA CoA -SH HS– 4

CH=H4F + H2O

H4F

H2O

H

Serine

H2N

COOH

Acetate– 5 COOH

CH=H4F CO2

H2N

C

H

CH2 SH Cysteine

Figure 3.5  Serine-glycine family amino acids biosynthetic pathways. Serine (Ser) is formed using: 1, 3-phosphoglycerate dehydrogenase; 2, 3-phosphserine aminotransferase; 3, 3-phosphoserine phosphatase. Synthesis of cystenin (Cys) requires: 4, serine transcetylase; 5, O-acetylserine sulfhrylase. Glycine (Gly) is formed using 6, serine hydroxymethyltransferase and decomposed by 7, glycine cleavage enzymes.

other hand in plants and microorganisms, since cysteine is synthesized by direct incorporation of HS-, cysteine serves as the universal sulfur carrier from which most of the sulfur containing metabolites incorporate sulfur directly or indirectly. Glycine synthesis is performed by the enzyme serine hydroxymethyltransferase where tetrahydrofolate is used to cleave a methyl group forming methyl-H4F and water. Enzyme expression in this pathway is repressed by several products of C1 metabolism, i.e., serine, glycine, methionine, purine, and thyamine.

3.2.4  Aromatic Nonpolar Amino Acids Erythrose 4-phosphate and PEP are the central precursor metabolites for the formation of the aromatic ring in the amino acids phenylalanine, tyrosine, and tryptophan (Figure 3.6). The C3 side-chain is also derived from PEP, or in the case of tryptophan from serine. In general, the biosynthesis of all three aromatic amino acids follows the shikimate pathway. It begins with the condensation of erythrose 4-phosphate and PEP into 3-deoxy-D-arabino-heptulosonate-7-phosphate (DAHP) catalyzed by three isozymes of DAHP synthase where each form of DAHP synthase is inhibited by a corresponding aromatic amino acid. Through the action of dehydroquinate synthase, DAHP is cyclized into 3-dehydroquinate,

3-11

Biosynthesis of Cellular Building Blocks: The Prerequisites of Life

Phosphoenolpyruvate (PEP)

Pi

CH2O P O OH

Pi

OH

OH

Erythrose 4-phosphate

OH

2

COOH OH 3-Deoxy-D-arabinoheptulosonate 7-phosphate (DHAP) Pi HO

1a, b, c

O

COOH OH 3-Dehydroquinate PEP 3-7

COOH O

Trp

–

HOOC

CH2

C

Phe

CH2

COOH Chorismate Glutamine 8 Pyruvate

– 13 a

O

C

OH

HO

Prephenate 14 b CO2, H2O

11 CO2, H2O

C H

N

CH

15

HO 12

HO NH2 H

O

N

O

C

C

COOH

Glutamate

P

CH2

15

Serine Glyceraldehyde 3-phosphate

C H

H

H Phenylpyruvate

OH CH

H H Tyrosine

COOH

PPi

Glutamate

C

COOH

10

H

H 4-Hydroxyphenylpyruvate

HOOC

HO PRPP

9 2 [H], CO2

C

O CH2 C

NH2 Anthranilate

13 b

HOOC

14 a

COOH

HOOC Prephenate

O

–

Tyr

NH2

H

NH2

C

C

COOH

H H Phenylalanine

CH CH2 COOH Tryptophan

Figure 3.6  Aromatic amino acids biosynthetic pathways. Common pathway is the shikimate pathway: 1a/b/c, 3-deoxy-D-arabino-heptulosonate-7-phosphate (DHAP) synthases; 2, dehydroquinate synthase; 3, dehydroquinate dehydratase; 4, shikimate dehydrogenase; 5, shikimate kinases; 6, 5-enoylpyruvoylshikimate-3-phsophate synthase; 7, chorismate synthase. Tryptophan (Trp) branch: 8, anthranilate synthase; 9, anthranilate-phosphoribosyl transferase; 10, phosphoribosyl-anthranilate isomerase; 11, indoleglycerolphosphate synthetase; 12, tryptophan synthase. Tyrosine (Tyr) branch: 13a/b, chorismate mutase/prephante dehydrogenase (Tyr-bifunctional enzyme). Phenylalanine (Phe) branch: 14a/b, chorismate mutase/prephante dehydrogenase (Phe-bifunctional enzyme); 15, tyrosine aminotransferase (used in both Tyr and Phe branches).

3-12

Cellular Metabolism

and converted into chorismate through a series of reactions catalyzed by dehydroquinate dehydratase, ­shikimate dehydrogenase, shikimate kinase, 5-enoylpyruvoylshikimate-3-phosphate synthase, and chorismate synthase. Conversion of chorismate is the first committed step towards any one of the three branching pathways leading to the aromatic amino acids. For the path leading to tyrosine, chorismate is converted into prephenate, and subsequently 4-hydroxyphenylpyruvate by a bifunctional enzyme chorismate mutase/ prephenate dehydrogenase using an intramolecular group transfer. Next, the incorporation of an amino group from glutamine by the enzyme tyrosine aminotransferase, results in the production of tyrosine. Similar to the initiation process of tyrosine biosynthesis, chorismate is converted into prephenate by chorismate mutase for biosynthesis of phenylalanine, but subsequently into phenylpyruvate by prephenate dehydratase. Finally, incorporation of an amine group from glutamate to phenylpyruvate by tyrosine aminotransferase results in phenylalanine formation. Tryptophan biosynthesis on the other hand starts with the formation of anthranilate from chorismate catalyzed by anthranilate synthase through the introduction of an amine group and a release of pyruvate. A series of reactions catalyzed by anthranilate-phosphoribosyl transferase, phosphoribosyl-anthranilate isomerase, indoleglycerolphosphate synthetase, and tryptophan synthase finally leads to the biosynthesis of tryptophan.

3.2.5  Histidine Biosynthesis Histidine is synthesized through a unique pathway involving ten biochemical steps, starting with an unusual condensation of ATP and 5-phosphoribosyl 1-pyrophosphate (Figure 3.7). 5-phosphoribosyl 1-pyrophosphate (PRPP) forms the C3 side-chain and the two adjacent carbon atoms −C = CH− of the heterocyclic ring. The purine base of ATP contributes to the atoms of the −N = CH− group. Then the conjugation of ATP with PRPP, followed by the release of pyrophosphate and incorporation of a water molecule, results in the synthesis of an intermediate in which the ribose ring opens up through the catalysis of phosphoribosylformimino-5-aminoimidazole carboxamide ribonucleotide isomerase. Subsequently an amide nitrogen is transferred from glutamine, followed by cleavage and ring closure to synthesize imidazole glycerol phosphate, with the other product, 5-aminoimidazole-4-carboxamide ribonucleotide, acting as an intermediate for the synthesis of purine (see Section 3.2.1). Next, a dehydration reaction which is catalyzed by histidinol-phosphate phosphatase results in the synthesis of imidazole acetol phosphate. Further transamination, from which the nitrogen group is derived from glutamate, as well as dephosphorylation, and dehydrogenation yields histidine. In enteric bacteria, the genes encoding the histidine biosynthetic enzymes form an operon assembly, arranged in the order of the reactions in the pathway.

3.3 Nucleotides as Building Blocks 3.3.1  Structure and Organization of Nucleotides The organizational blueprint for cellular machinery is made up of nucleotides. When these monomers are chained together to form the DNA and RNA strands, they act as the repositories and transmitters of genetic information for every cell, tissue, and organism. The monomer units are referred to as nucleotides, and are classified into the two main structures of purines and pyrimidines. Nucleotides are also known to serve additional functions throughout the cell, involving a number of different metabolic objectives and account for up to 20% of the cells dry weight in prokaryotes. 3.3.1.1  Primary Structure of the Nucleic Acids Nucleotides are composed of three main components: (1) the main ribose or deoxyribose sugar, (2) the phosphate linker, and (3) the nucleobase. When the ribose sugar is in the unphosphorylated state at the 5´ carbon and contains only a nucleobase attached the 1´ carbon, the molecules are known as nucleosides.

His OH OH

NH2 O

N

N

n O

N H

O

CH2

N N

8

H 2O

CH

Glutamate

C H2

O

N

N

N

Pi

9

5

NH2

3

H2O

Histidinol

HO CH2 CH CH2

AICAR

2

PPi

Ribose 5’ -P

2 [H]

N

H2C

P O

H2C

P O

10

O H C

HOOC

O

CH2

OH HN

2 [H]

N

HO

HN

OH OH

O

O H C

4

N

N

Histidine

N N

Ribose 5’ -P

Ribose 5’ -P

N

N

N

CH CH2

NH2

NH2

N

NH2

Figure 3.7  Histidine biosynthesis. Histidine (His) is synthesized via a complex series of reactions. Numbered enzymes are: 1, phosphoribosyl-ATP pyrophosphorylase; 2, phosphoribosyl-ATP pyrophosphohydroylase; 3, phosphoribosyl-AMP cyclohydrolase; 4, phos-phoribosylformimino-5-aminoimidazole carboxamide ribonucleotide isomerase; 5, phosphor-ribosylformimino-5-aminoimidazole carboxamide ribonucleotide; 6, cyclase; 7, imidazole-glycerolphosphate dehydratase; 8, histidinol-phosphate transaminase; 9, histidinol-phosphate phosphatase (as a bifunctional enzyme with 7 or separate); 10, histidinol dehydrogenase.

P O CH2 C

ke to ni za tio

H2O

6

OH OH

P O CH2 CH CH

OH OH Imidazole glycerol phosphate

7

H2C

P O

Resynthesis of purine ring

Glutamine

1

–

PPi

P O CH2 CH CH

(ATP) Adenosine 5’ -triphosphate

(PRPP) 5-Phosphoribosyl 1-pyrophosphate

N

Biosynthesis of Cellular Building Blocks: The Prerequisites of Life 3-13

3-14

Cellular Metabolism

On the other hand when the monomer ribose sugar is phosphorylated at the 5´ carbon and has a base attached it is termed a nucleotide. The nucleobase is always connected to the 1´ position using a glycosidic bond while the ribose sugars are connected from the 3´–5´ phosphodiester bond of successive ribose sugars (Figure 3.8). These bonds form the backbone of the nucleic acid molecule. The nucleotide chains are arranged in one direction which is established by the bonds formed between the 5´-phosphate and the 3´-hydroxl group. The specific nucleotide sequence, referred to as the primary structure of the DNA is the storage for the genetic information for protein synthesis and is read from the 5´ to the 3´ end. 3.3.1.3 The Basic Components of Nucleotides Nucleosides are classified according to their nucleobases into purines (nine carbon ring molecules), which include adenosine (A) and guanosine (G), and pyrimidines (six carbon ring molecules), which are cytidine (C), uridine (U), and thymidine (T). Instead of thymidine, uridine (U) is used for RNA assembly (Figure 3.9). Moreover, RNA is also assembled from the ribose sugar molecules, and not from the deoxyribose form used for DNA formation. The labeling of the atoms in ribose generally includes a prime (′) to differentiate between the labeling of the base carbons. As mentioned earlier, DNA links the two strands through hydrogen bonds of the complementary pairing nucleotides A-T/U and G-C.

(O) P O– O

5´

Base

Base N

CH2

N

O

O (O) P O–

CH2 – O

O H

3´

H 2´

OH

2´

O –O P (O)

5´ N

N

O

OH

Base

Base

O

3´

2´

5´

CH2

O

H

2´

OH

CH2 – O

O

Hydrogen bonding

H

3´

–O P (O)

5´

3´ OH

O

Figure 3.8  Nucleotide backbone connections. The phosphodiester and glycosidic bonds are evident in the connection of consecutive nucleotides. Hydrogen bond between strands causes a rotation of the strands resulting in the helix structure so often depicted. 3′ and 5′ locations have been identified. (a)

CO2

6 Aspartate

1N

5

2

3N

4

N7

N9

HC Glutamine

Ribose 5’ -P Glutamine

OH

Carbamoyl phosphate 8

HOCFH4

(b)

Glycine

OH

6 1

FH4 O

2

Aspartate 5

N N3

4

Ribose 5’ -P

Figure 3.9  Origin of each atom making up the nucleotide precursors. (a) IMP, the first synthesized purine, and (b) UMP, the first synthesized pyrimidine.

3-15

Biosynthesis of Cellular Building Blocks: The Prerequisites of Life

The phosphate linker can be a monophosphate, diphosphate, or triphosphate group attached to the 5´ carbon of the ribose. The monophosphate forms of AMP, CMP, GMP, UMP, and TMP molecules are used as energy sources through cellular metabolism. Because the nucleoside monophosphates (NMPs) and diphosphates (NDPs) are used as energy sources and also help to carry phosphates around the cell, the have a very high rate of formation. Both are also repeatedly phosphorylated with ATP by nucleotide specific NMP and NDP kinases that can act on both the deoxyribose and ribose forms.

3.3.2  Biosynthesis of Nucleotides Both purine and pyrimidine biosynthesis are feedback inhibited to synchronize physiological demand within the cell. Production is regulated by intracellular mechanisms that sense and control the availability of nucleotide triphosphates (NTPs) as they accumulate and dissipate during growth and tissue regeneration. In general, regulations of nucleotide synthesis are complex, and further information can be found elsewhere for detailed information is beyond the scope of this chapter. The 2-deoxyribonucleotides are catalyzed by ribonucleotide reductase which reduces a ribonucleotide into the deoxyribonucleotide. The nucleoside triphosphates are all, with the exception of CTP, synthesized from the corresponding monophosphate forms. The pentose portion of both purines and pyrimidines are derived from PRPP, but at different points in the synthesis. In addition, NDP-sugars can also be synthesized directly from a few central precursor metabolites (Figure 3.10). Purines add a number of different groups to PRPP and eventually from the heterocyclic ring while in pyrimidine synthesis the PRPP is introduced after the ring has already been formed. 3.3.2.1  de novo Purine Synthesis In humans, a limited amount of nucleic acids derived from diets are incorporated into the cells and thus must be synthesized de novo. There are two major routes of purine biosynthesis; the first is synthesis from amphibolic intermediates, more commonly known as synthesis de novo and the second is through the use of salvage pathways (see Section 3.2.4). In de novo synthesis, the active ribose 5-phosphate, PRPP, is converted into inosine monophosphate (IMP) through a series of ten enzymatic reactions where functional groups are added in a stepwise manner. IMP is then used to form the final purines adenosine monophosphate (AMP) or guanosine monophosphate (GMP) through the inclusion of the respective base (Figure 3.11). Fructose 6-phosphate

Mannose 6-phosphate

Glucose 6-phosphate

Glutamine

Glucosamine 6-phosphate Acetyl -CoA

Mannose 1-phosphate

Glucose 1-phosphate

N-Acetylglucosamine 1-phosphate

NTP

NTP

NTP

PPi

PPi

PPi

NDP-mannose

NDP-glucose

NDP-N-acetylglucosamine

Figure 3.10  Activation pathways for hexose 1-phosphates from central precursor metabolites. While some can be activated by nucleotidyl transfer from the corresponding triphosphate, most other activation mechanisms occur by using a diphosphate form. UDP-galactose can be formed directly from galactose by UDP activation.

3-16

Cellular Metabolism

ATP

Glutamine PPi 5-Phosphoribosyl 1-pyrophosphate (PRPP)

– –

NH2

ADP + Pi

Ribose 5’ -P

1

C O

2

C

Ribose 5’ -P -Glycineamide

AMP GMP 5‘ -phosphoribosyl

CH=H4F ADP + Pi

N N

H N

ATP

5

C

NH

HN

Ribose 5’ -P

ADP + Pi

H

O

3

ATP Glutamine

H N

4

NH

O

Ribose 5’ -P

-5-Aminoimidazole

C

H

O

Ribose 5’ -P

-N-Formylglycineamidine

-N-Formylglycineamide

6

CO2

Aspartate

HOOC

ATP

N

ADP + Pi

O

Fumarate H2N

N

H2N

7

8

C

Ribose 5’ -P

9 Ribose 5’ -P

-4-Carboxamide5-aminoimidazole

2 [H] H2O

N

N

(IMP) Inosine 5’ -monophosphate

12

H2N

N

N H

N Ribose 5’ -P

H 2O

Aspartate GTP GDP + Pi 13 8

Fumarate

NH2 N

N

N

-4-Carboxamide5-formaminoimidazole

OH

Glutamine ATP ADP + Pi

N Ribose 5’ -P

(XMP) Xanthosine 5’ -monophosphate

C

10 N

N

11

Ribose 5’-P

H C

O

OH N

N

H2N

N

OH N

O

OHC-H4F

N

H2N

-5-Aminoimidazole4-carboxylic acid (AICAR)

HO

NH

Glycine

-Amine

H2N

NH2

N Ribose 5’ -P

(GMP) Guanosine 5’ -monophosphate

N

N N

N Ribose 5’ -P

(AMP) Adenosine 5’ -monophosphate

Figure 3.11  Purine synthesis from precursor 5-Phosphoribosyl 1-pyrophosphate (PRPP). Using a ten step irreversible pathway IMP is formed from: 1, PRPP amidotransferase; 2, phosphoribosyl-glycineamide synthetase; 3, phosphoribosyl-glycineamide formyltransferase; 4, phosphoribosyl-formylglycineamidine synthetase; 5, phosphoribosyl-aminoimidazole synthetase; 6, phosphoribosyl-aminoimidazole carboxylase; 7, phosphoribosyl-aminoimidazole succinocarboxamide synthetase; 8, adenylosuccinate lyase (bifunctional enzyme); 9, phosphoribosyl-aminoimidazole carboxamide formyltransferase; 10, IMP cyclohydrolase; GMP is then formed using 11, IMP dehydrogenase and 12, GMP synthetase while AMP is synthesized using 13, adenylosuccinate synthetase and the bifunctional enzyme.

Biosynthesis of Cellular Building Blocks: The Prerequisites of Life

3-17

To begin the purine synthesis de novo, an imidazole ring is formed by the introduction of glutamine and glycine in five reaction steps which is then followed by the completion and closure of the pyrimidine ring using aspartate. Overall, several different functional groups are added from glycine molecules, amino groups from glutamine and aspartate, single carbons from methenyl or formyl containing compounds and a final carbon from carbon dioxide. The first step involves the formation of PRPP by PRPP synthase using a free ribose 5-phosphate and ATP. The 1′ carbon is then aminated by PRPP glutamyl amidotransferase using glutamine and water followed by the addition of glycine to the recently added NH3+ . Glycine contributes the C4 and C5 carbons as well as the N7 nitrogen of the imidazole ring. Reaction with tertrahydrofolate adds a carbon to the C8 which is acted on by a series of synthetases (VI and VII) adding the N3 in the pyrimidine ring and closing the imidazole ring. The subsequent five reactions, beginning with a carboxylation, complete a purine structure forming IMP. AMP is then formed by aminating IMP at the C6 position by adenylosuccinase. In forming GMP, the C2 position of IMP is first oxidized to form xanthine monophosphate (XMP, nucleobase of xanthine) and then aminated at the same C2 position to create GMP. Later phosphorylation leads to the diphosphates ADP and GDP and then to the triphosphate forms ATP and GTP, respectively. Both prokaryotic and eukaryotic synthesis de novo steps are the same but there are major differences in the functionality of the enzymes used to catalyze the reaction steps. While in prokaryotic cells each step is catalyzed by its own unique enzyme, in purine producing eukaryotic cells the enzymes contain multiple catalytic activities facilitating the successive conversions through adjacent catalytic sites. For instance the addition of glycine, the C8 position and closing of the imidazole ring are performed at different catalytic sites on the same functional enzyme in human liver cells. Since a number of precursor metabolites (glycine, glutamine, ATP, aspartate, and tetrahydrafolate) are incorporated into nucleotide biosynthesis, it is vital that the metabolism of nucleotides is tightly regulated to not induce further cell stress. For example, PRPP synthase is sensitive to the concentration of AMP, ADP, GMP, and GDP where high concentrations result in feedback inhibition of the enzyme. AMP also limits its own formation by inhibition of the adenylosuccinate synthase enzyme, as does GMP with its own inhibition of IMP dehydrogenase. There is also cross regulation that occurs for the generation of GMP and AMP in IMP metabolism where a decrease in synthesis of one occurs when the other is in a state of deficiency. 3.3.2.2  Stepwise Production of Pyrimidines The de novo biosynthetic pathway of pyrimidines occurs in a disjointed manner since the active ribose phosphate is not incorporated until after the pyrimidine ring is formed. Pyrimidine synthesis is regulated through allosteric control of the three enzymes CTP synthase, carbamoyl phosphate synthase, and aspartate carbamoyltransferase (Figure 3.12). To start the biosynthetic pathway, the enzyme carbamoyl phosphate synthase (CPS) condenses aspartate and carbamoyl phosphate to form carbamoyl aspartic acid (CAA). In eukaryotic species, there are two forms of the CPS enzyme, mitochondrial (Type I) and cytosolic (Type II) protein. Only CPS II is responsible for the biosynthesis of pyrimidines from its independent pool of carbamoyl phosphate. In prokaryotes, there also exists multiple forms of the CPS enzyme though not typically classified as Type I or II and all are functionally active. After formation of CAA, the ring closure occurs to from dihydroorotic acid which then is converted to orotic acid by dihydroorotase dehydrogenase. Next, orotic acid is attached to PRPP by orotate phosphoribosyltransferease and then decarboxylated to from UMP. Before forming either TMP or CTP, uridine monophosphate is always phosphorylated to UDP. Another phosphorylation generates UTP before being converted by CTP synthase using ATP and glutamine to create CTP. Alternatively UDP is acted on by ribonucleotide reductase to remove the formed deoxyuridine diphosphate (dUDP) which after a dephosphoylation is converted to dTMP by thymidylate synthase, then oxidized to form the final pyrimidine building block TMP. 3.3.2.3  Formation of 2´-Deoxyribonucleotides In order to form 2-deoxyribonucleotides, ribonucleotides must be synthesized first through the steps outlined in the previous sections, and then after phosphorylation to the diphosphate and triphosphate

3-18

Cellular Metabolism NH2

O C

HO

CH2

CH

COOH

OH

Pi

Aspartate

C O O

O O

H 2N

H2N

1

P

CH2

N

CH N COOH O H Carbamoyl aspartate

OH OH

OH

H2O

C

2

C

O

C

CH

N H

COOH

4,5 -Hihydroorotate

Carbamoyl phosphate

3 2 [H]

OH

OH CO2

PPi

OH

PRPP

N

N O

CH2

N 5

N

O

4

COOH

N

HO

Ribose 5’ -P

Ribose 5’ -P (UMP) Uridine monophosphate

N

COOH

Orotate

Orotidine monophosphate

ATP 6

ADP OH

OH ATP ADP

N O

N

7 Ribose 5’ -PP

(UDP) Uridine diphosphate

Glutamine ATP ADP + Pi

N O

NH2

N

8 Ribose 5’ -PPP

(UTP) Uridine triphophate

N O

N Ribose 5’ -PPP (CTP) Cytidine triphosphate

Figure 3.12  Synthesis of pyrimidine nucleotides via orotate. The enzymes used are: 1, aspartate transcarbamoylase; 2, dihydroorotase; 3, dihydroorotate dehydrogenase; 4, orotate phosphoribosyltransferase; 5, orotidine 5-phosphate decarboxylase; 6, nucleoside-momophosphate kinase; 7, nucleoside-diphosphate kinase; 8, CTP synthase.

forms, ribonucleotide reductase can catalyze the reduction reaction to the deoxy- forms. This highlyevolved and highly regulated enzyme acts using a radical donor mechanism involving a coenzyme B12 ion, an iron ion, or a noniron dependent enzyme. The basic mechanism for ribonucleotide diphosphates involves the donation of an electron, generally by reduced thioredoxin, which is then later oxidized back to its reduced state by the cofactor NADPH. The deoxyribonucleoside diphosphates are then later phosphorylated by ATP to triphosphates. The reduction products, deoxyuridine triphosphate (UTP) and deoxyuridine diphosphate (UDP) are not the DNA building blocks but the main enabler for deoxynucleotides. UTP and UDP are dephosphorylated and reduced into dUMP by either the enzyme dUTP pyrophosphatase or dUDP phosphatase, respectively. dUMP is then converted into the deoxythymidine monophosphate (dTMP) through the action of the enzyme thymidylate synthase. In the final reaction to form dTMP, methylene

Biosynthesis of Cellular Building Blocks: The Prerequisites of Life

3-19

tetrahydrofolate donates the methylene group that is simultaneously reduced to the methyl level by oxidizing the carrier tetrahydrofolate to dihydrofolate, later reducing itself back to tetrahydrofolate. dTTP is though to be formed through a different mechanism since production levels seam to be independent of activity through dUTP pyrophosphatase. 3.3.2.4  Production through Salvage Reactions As mentioned before, the regulation of de novo synthesis is very tightly controlled resulting in instances where the production levels can not meet cellular demands. As such, and especially in prokaryotes, there exists a collection of salvage enzymes to help sustain production levels in periods of high demand. Salvage reactions generally involve less energy than primary synthesis routes since the major pieces of the nucleotide molecules have already been formed, thus only requiring a few more steps and metabolic precursors to complete the nucleotide biosynthesis. The salvage enzymes in prokaryotes have the primary function of allowing nucleotide synthesis from free nucleosides and nucleobases that permeate into the cell while also recycling back signaling molecules, such as cAMP, and any nucleobases or nucleosides produced from nucleotide turnover. In addition, salvage pathways also provide carbon, energy, and nitrogen by deconstructing the dephosphorylated nucleotides that have entered the cell through periplasmic membranes and phosphatases. Salvage reactions occur by a number of mechanisms and variations; however, there are five typical mechanisms: (1) a free purine and PRPP form a purine 5′ mononucleotide with a pyrophosphate where two phosphoribosyl transferases combine the free purine to the one carbon position of PRPP. This is the most common method for recycling of nucleobases as it has an equilibrium strongly favoring the assembled nucleotide. (2) Nucleoside kinases transfer a phosphoryl group in a phosphorylation reaction from nucleotide triphosphates to form mononucleotides. This reaction does not occur in all bacteria species due to the lack of metabolic precursors needed in the reaction mechanism, but does occur in most mammalian cell lines. (3) Reversion between nucleobases and nucleosides can occur by nucleoside phosphorylases, which places a phosphate O-glycosidic linkage in place of the N-glycosidic linkage. (4) The N-glycosidic linkage of monophosphates can be hydrolyzed by NMP glycosylases for formation of dNMPs. (5) Since the nucleotide synthesis steps are not reversible, mechanisms for the interconversion of nucleobases exist, such as the conversion of adenine to hypoxanthine (the nucleobase for IMP) and deamination of GMP to create IMP. In general, salvage reactions play a major role in the replenishment of the nucleotide pool and are the major method of nucleotide generation in humans. More specifically, the human liver is the primary location used in generating purines and purine nucleosides used in the salvage pathways. This generation is critical for other tissues in the body as well. For instance, the human brain has been found to produce only small quantities of PRPP synthase, the enzyme responsible for synthesis of PRPP, and therefore it must rely on these exogenous purines from the liver. Similarly erythrocytes and polymorphonuclear leukocytes cannot synthesize 5-phosphoribosylamine, a precursor synthesized by the PRPP glutamyl amidotransferase.

3.3.3 Nucleotide Metabolic Conditions Abnormal nucleotide catabolism can result in a number of clinical disorders ranging from mild to fatal conditions in humans where the majority of nucleotide related diseases result from purine catabolism problems. Gout is a clinical condition caused by the excess accumulation of uric acid, a by product of purine degradation, and typically results in severe inflammation of the joints or similarly arthritis that is caused by a precipitation of sodium urate crystals. Readers can find a more information in the review by Nyhan [25]. The most severe of purine metabolism related diseases are Lesch–Nyhan syndrome and Severe Combined Immunodeficiency Disease (SCID). Lesch–Nyhan syndrome is caused by an inherited defect in the X chromosome which results in the loss of function in the HGPRT gene (encodes a nucleotide

3-20

Cellular Metabolism

salvage enzyme). In the most serious cases, patients not only have severe symptoms of gout, but a severe malfunction of the nervous system usually resulting in death before the age of 20 [24]. SCID is the resulting condition of a deficiency in the adenosine deaminase enzyme that converts adenosine to inosine and selectively leads to the destruction of B and T lymphocytes responsible for mounting the body’s immune response [2,3]. Often seen in cancer patients undergoing chemotherapy, conditions of purine deficiency occur due to treatment methods that inhibit the formation of tetrahydrofolate, a major precursor metabolite of purine synthesis. These treatments are designed to limit tumor growth in patients, however since the initial carbons used to form the purine rings are derived from tetrahydrofolate derivatives, these deficiency states result. Typically this is diagnosed by monitoring the levels of folic acid in the body, the metabolite responsible for the pool of tetrahydrofolate. While there are only a few disorders caused by pyrimidine metabolism, they are still prevalent in the population. The two main disorders are inherited disorders affecting the bifunctional enzyme catalyzing the UMP synthesis, orotate phosphoribsyl transferase and OMP decarboxylase. Deficiencies in these enzymes result in orotic aciduria causing retarded growth and conditions of hyporchromic erythrocytes and megaloblastic bone marrow. Both conditions are prevalent in children and lead to severe anemia [25].

3.4  Synthesis of Carbohydrates for Building Cells 3.4.1  Introduction to Carbohydrates Carbohydrates, also commonly referred to as saccharides, are found attached to all forms of natural molecules such as proteins, lipids, hormones, and antibiotics. As such, they are critically important building blocks for proper cell functioning by both providing energy for metabolism through oxidation and serving as structural components for a variety of cellular membranes. Understanding such roles in cellular function requires elucidation of the mechanisms by which interconversions between carbohydrates take place, the role they play as building block precursors as well as modes in which they are introduced to pathways by activation. While monosaccharides and their derivatives make up the majority of the carbohydrates found in cells, polysaccharides, oligosaccharides and several interlinked carbohydrate monomers are also highly prevalent. Originally the term carbohydrate was reserved to only those molecules containing the traditional Cn(H2O)n structure (hydrates of carbon), but now is also used to include molecules having similar physical properties such as the polyhydroyl aldehydes, alcohols, ketones, and acids. In plants, animals, and microorganisms, the carbohydrate glucose is one of chief importance since it is used as an energy source, primarily through the main oxidative pathway of glycolysis. Glucose is generated from a variety of metabolic routes including simple transport, oxidation, and reduction. For example in plants, glucose is derived from the energy-storage polymer starch produced during photosynthesis in which, water and carbon dioxide are converted into carbohydrates and oxygen with the use of sunlight as an energy source. The primary form synthesized by plants is D-glucose, which is utilized to generate plant polymers such as cellulose and even reconverted back to starch storage. In humans carbohydrates are primarily derived from plant fiber intake, which is broken down into water and energy. A similar process is used by most bacteria where glucose and other simple sugars are imported into the cell through various transport mechanisms. Carbohydrates are generally separated into two main groups: simple and complex sugars. Simple ­sugars are generally monosaccharides containing only the carbohydrate structure itself. This class of ­sugars includes mannose, fructose, and ribose along with their various derivatives. Monosaccharides are ­identifiable through the number of carbon atoms present (e.g., pentoses have five carbon atoms, where hexoses have six), the configuration of the hydroxyl group, and the position of the carbonyl group and are joined together by acetal-linkages to form oligosaccharides and polysaccharides. In many instances, monosaccharides, such as glucose, exist in different isomeric forms (D and L form) where the D sugars are the most naturally abundant. As such, most enzymes in mammals are selective to the D sugars.

Biosynthesis of Cellular Building Blocks: The Prerequisites of Life

3-21

The most stable forms of monosaccharides occur with ring structures similar to pyran and furan and may also exhibit alpha and beta anomers, especially when the compounds are exposed to different environmental conditions such as pH. Additionally, several other varieties can occur in sugar molecules, which include epimers, different configurations of hydroxyl groups, aldose-ketose isomers, and structural conformations. Because of the important modifying influence these different isomeric forms take in addition to their attachment to many lipid and benzenoid structures, numerous enzymes called glycosidases have evolved for the attachment and removal of sugars. The advanced simple sugar disaccharides such as maltose, sucrose and lactose are also critical carbohydrates prevalent in many organisms. Many humans suffer from the inability to metabolize lactose, the disaccharide most prevalent in milk; this is something that leads to diarrhea, flatulence and other serious metabolic conditions. Maltose, or malt sugar, is used in the hydrolysis of starch, which itself is a carbohydrate polymer. Many polysaccharides have a major physiological role in cellular function, specifically the homopolymer starch, the storage molecule glycogen and the related molecules glycol-aminoglycans and glycoproteins. Complex carbohydrates make up a broad spectrum of molecules where carbohydrate molecules are covalently bonded to a lipid, protein, or other larger biomolecular structure, examples of which include glycosides, glycolipids, antibiotics, glycoproteins and proteoglycans, peptidoglycans, and lipopolysaccharides. Peptidoglycans are the highly branched macromolecules in cell walls of plants providing membrane strength made up of linear polysaccharide strands that are cross-linked by oligopeptide units. Gram-negative bacteria use the highly complex lipopolysaccharides in their outer cell envelope where these molecules sometimes serve as receptors to detect bacteriophages intent on causing infection. One of the most important groups of complex carbohydrates are antibiotics, since they not only serve as simple solutions to some serious health problems but are essential tools used in the latest research biochemistry techniques. This group of compounds is excreted by microorganisms to inhibit the growth of other organisms generally through the interference of DNA transcription, RNA translation, or protein synthesis. A sub class called nucleoside antibiotics generally contain a normal base attached to a carbohydrate other than the D-ribufuranose sugar. The first nucleoside antibiotic discovered was puromycin, a potent antibiotic causing premature chain release during translational chain elongation by mimicking the 3′-end of tRNA. Aromatic antibiotics contain one or two sugars attached to an aromatic molecule having a polycyclical character, for example the cancer cell killing drug daunomycin [30]. Another potent antibiotic is erythromycin, where the sugar moiety is attached to a larger lactone ring which contains the sugar activation sites. Interestingly, the macrolide antibiotic will lose its function completely when the sugar moiety is removed from the lactone ring. Because of its natural chemotherapeutic properties as well as being a stable molecule for biochemistry protocols, the most widely used antibiotic today is streptomycin. It is a member of the aminoglycoside antibiotics that has an aminocyclotol attached to amino sugars which are themselves linked to other sugars [28]. Glycolipids occupy a minor part of the overall molecular structure and commonly occur with lipids associated with outer cell surfaces. By inhibiting biological activity of toxins, glycolipids aid in molecular transport across cell membranes and are also known to be involved with the biosynthesis of glycoproteins and proteoglycans. Glycoproteins and proteoglycans are similar in that both are proteins containing chains of carbohydrates linked by N- or O-glycosidic bonds but differ in the configuration of attached carbohydrates to the chain. For clycoproteins, the carbohydrates are a short, branching of complex monomer units where proteoglycans are long, simple linear strands of carbohydrate monomers. Glycoproteins are usually ­15–20 repeating monosaccharide units making up the chains typically consisting of D-glucose, D-galactose, deoxy-D-galactose, D-mannose, and L-fucose. The saccharide chains of proteoglycans are approximately 100 monomers in length with disaccharide repeating units usually consisting of a hexosamine and a hexuronic acid with a sulfate group attached. The addition of the sulfate makes these molecules highly charged and thus naturally very reactive species. Nevertheless, both of these large molecules are included in structures such of hormones, enzymes, collagens among others and are evenly dispersed through cells of higher animals, plants, and microorganisms.

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Cellular Metabolism

3.4.2  Interconversion of Carbohydrates The monosaccharides, as principle molecules of metabolism and cell function, are synthesized through a variety of routes by an array of enzymes responsible for their physiological functional levels. Sugars containing three and six carbon molecules are generated through glycolysis/gluconeogenesis; however the other central precursor sugars having four and five carbons are synthesized through separate metabolic channels. The four major routes leading to these carbohydrate building blocks are: (1) The oxidative pentose-phosphate pathway, which is essential to all organisms for the interconversion of many sugar phosphates producing the central precursor metabolites for the synthesis of some amino acids, nucleotides, and other important building blocks. Most bacteria are capable of using the main carbohydrate degradation pathway as well as the oxidative pentose-phosphate pathway or the Entner–Doudoroff pathway. (2) Formation of NDP-sugars from fructose 6-phosphate in gluconeogenesis. (3) Transfer ­reactions used to activate the (NDP)-sugars. Such reactions include glycosyl transfer, pyrophosphoryl transfer, or nucleotidyl transfer. For example, the activation of sugar 1-phosphates with NTP via nucleotidyl transfer or the ribose 5-phosphate activation to PRPP, which itself acts as a mode of synthesis for nucleotides. (4) Synthesis of activated sugars. Sugars must be metabolized further into usable building blocks through the formation of glycosidic bonds with various acceptor groups, or other activated forms. 3.4.2.1  Pentose Phosphate Pathway The pentose phosphate pathway’s primary function is to efficiently replenish the pool of NADPH but also serves to keep the carbon flow towards the primary metabolic targets in the end of glycolysis (Figure 3.13). This pathway is also the main mode of interconversion from hexose 6-phosphates into triose 3-phosphates and other carbon phosphates. Pathway initiation begins with glucose 6-phosphate (G6P) converted into 2-dehydrogluconate by the reversible action of G6P dehydrogenase, an enzyme inhibited by high levels of NADPH. In a competing reaction, G6P is converted to fructose 6-phosphate by phosphoglucose isomerase (PGI), an important enzyme in glycolysis universal to plants, animals, NADPH NADP+ + H+ Glucose 6-phosphate

Gluconolactone 6-phosphate

1

NADPH + H+

NADP+

H2O

6-Phosphogluconate

2

3

Ribulose 5-phosphate 4

Fructose 6-phosphate

Glyceraldehyde 3-phosphate

6 C2

Erythrose 4-phosphate Xylulose 5-phosphate

Glycolysis

Xylulose 5-phosphate

Glyceraldehyde 3-phosphate C3

5

7

C2 6 Sedoheptulose 7-phosphate

To nucleotide synthesis

Ribose 5-Phosphate ATP 8 AMP PRPP

Figure 3.13  Pentose phosphate pathway’s oxidative routes for NADPH and pentose formation. Three hexose molecules cycle through the action of: 1, glucose 6-phosphate dehydrogenase; 2, lactonase; 3, 6-phosphogluconate dehydrogenase; 4, ribose 5-phosphate isomerase (can proceed in the reverse direction); 5, ribulose 5-phosphate 3-epimerase; 6, transketolase; 7, transaldolase; 8, PRPP synthetase.

Biosynthesis of Cellular Building Blocks: The Prerequisites of Life

3-23

and bacteria. PGI strongly favors the formation of fructose 6-phosphate under both aerobic and anaerobic conditions in E. coli [34], unless a cellular demand exists for NADH or NADPH. E. coli mutants lacking the gene for PGI grow more slowly on glucose and other sugars since the pentose phosphate and Entner-Doudoroff pathways are left to make up the lack of metabolites down stream of PGI [14]. Continuing the pentose phosphate pathway, 2-dehydrogluconate undergoes two irreversible steps leading to the formation of ribulose 5-phosphate through oxidation and the generation of NADPH. Ribulose 5-phosphate is then processed by phosphoribulose epimerase to form xylulose 5-phosphate or by phosphopento isomerase to form ribose 5-phosphate. These two precursors are then interconverted to form other sugar phosphates, such as erythrose 4-phosphate and sedoheptulose 7-phosphate, that play important roles in phosphorylation, coenzymes, and in transport and storage. Xylulose 5-phosphate and ribose 5-phosphate are converted by transketolase (TK) to form the metabolites glyceraldehyde 3-phosphate and sedoheptulose 7-phosphate. Transaldolase (TA) then converts these to form fructose 6-phosphate and erythrose 4-phosphate. TK exchanges a carbon between erythrose 4-phosphate and xylulose 5-phosphate to form fructose 6-phosphate and glyceraldehyde 3-phosphate. The yeast TK has been investigated for a number of industrial synthesis purposes and has been shown to have a high level of selectivity for diasteric molecules, those with the C2 steroiochemistry [8]. On the other hand, no wide spread study has yet been performed on the substrate specificity exhibited by TA. The complex network of the pentose phosphate pathway allows for the generation of these glycolysis intermediates, enabling bacteria to grow on a variety of different carbon sources thus allowing a high degree of metabolic flexibility. In particular, when glycolysis is inhibited even under cell states of high NADPH, the pentose phosphate interconversions allow for the synthesis of fructose 6-phosphate, and the excess NADPH can be converted to energy through oxidative phosphorylation routes. 3.4.2.2 NDP-Sugars as Building Blocks NDP-sugars are formed using central precursors through a few simple enzymatic reactions (see Figure 3.10) beginning with the isomerization of ribose to ribulose (as seen mentioned above). Isomerases ­interconverting an aldo group to a keto group in the synthesis of carbohydrates have been observed to exhibit substrate specific reactivity towards hexoses, pentoses, and nonphosphorylated pentoses. The isomerization of esters has long been considered an important part of glycolysis and as such is recognized as vital to pentose metabolism. Mutases are then employed to manipulate the phosphate groups, for example that of 6-phosphohexoses which is relocated to the C1 carbon to form 1-phosphohexoses. Alternatively, formation of amino sugars is done by transamination of fructose 6-phosphate to glucosamine 6-phosphate where acetyl-CoA is used as an acyl donor for the formation of the N-acetylated sugar. The sugar 1-phosphates are then converted to activate sugars in an irreversible manner. Activation can be accomplished via three methods: glycosyl transfer, pyrophosphoryl transfer, or nucleotidyl transfer. Glycosyl transfer is primarily used in the synthesis of complex carbohydrates. In pyrophosphyl transfer, a pyrophosphate group is transferred to pentose 5-phosphate from ATP, which then yields the activated pentose 5-phosphate and AMP. The enzyme PRPP synthetase acts on ribose 5-phosphate to form PRPP through this mechanism. Nucleotide transfer involves the activation of ­a ldohexoses by adding the nucleotide to the sugar releasing pyrophosphate in the process. An example of this is the conversion of glucose 1-phosphate to UDP-glucose by the NTP-hexose-1-phosphate nucleotidyl transferase (a similar reaction is used to create NDP-mannose from mannose 1-phosphate). It is important to note that the transferases used are specific to the sugar moiety and the type of NTP involved thus predefining the location of the sugar and conveniently are strictly controlled by feedback inhibition. The reaction equilibrium is such that the reaction favors the formation of the NDP-sugars creating excess free energy for the synthesis of nucleotides and other sugar-containing polymers. The activated sugars can then be used as building blocks or further converted to other activated sugars. NDP-sugars are incorporated during a carbohydrate polymerization process which forms critical polymers used throughout the cell such as: (a) disaccharides and polysaccharides including glycogen, (b) lipid diphosphate sugar intermediates, and (c) glycosides. NDP-sugars can also play functional roles

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Cellular Metabolism

in the synthesis of complex carbohydrates. However, before polymerization occurs, the NDP-sugars must be activated, modified for specific function and then transferred to the appropriate receptors. In prokaryotes, NDP-sugars are the major precursors leading to the formation of most of the activated building blocks used in the formation of carbohydrate-containing polymers, specifically acting as the glycosyl donors. Additionally, they are known to act as regulatory mediators during the polymerization process. Since the nucleotidyl transferases are specific to the NTP and sugar molecules, the transferases act as built-in metabolic regulators by controlling the channeling of carbon into separate pathways leading to different sugar-containing polymers. Frequent interconversion between NDP-sugars also takes place by a variety of alternative mechanisms. UDP-glucose can undergo oxidation to form a ketosugar and then is reduced to form uronic acids with the retention or inversion of hydroxyl group configurations. The oxidation is performed by NDP-sugar dehydrogenases using NAD + as an electron acceptor. UDP-glucose can also undergo epimerization to form UDP-galactose via UDP-glucose 4-epimerase or to form UDP-mannose by UDP-glucose 2-epimerase. Finally, the deoxy-NTP sugars can undergo a series of reductions, oxidations, and epimerizations to form the 6-deoxy hexoses such as dTDP-6-deoxy L-talose and dTDP-L-rhamnose. Another way in which the flow of carbohydrate is regulated is during the transport of NDP-activated sugars across membranes where the pathway used is dictated by the type of sugar. The transport of NDPsugars across hydrophobic lipid layer of the cytoplasmic membrane is regulated by lipid carries that attach carrier-specific sugars to them for transit across the membrane. These carriers are highly important since all the NDP-activated sugars making up extracellular polymers are water soluble, whereas the carries are not, thus stabilizing the polymer. For example, the carrier undecaprenyl phophate aids in the synthesis of peptidoglycan and other glycans whereas dolichol phoaphates are utilized during synthesis of glycoproteins.

3.4.3 Other Import Enzymes of Carbohydrate Synthesis 3.4.3.1  Critical Kinases Kinases, which transfer phosphates from ATP to carbon structures, are responsible for the variety of carbohydrates found in microorganisms. The formation of kinases, either constitutively or in response to added inducers, directly leads to the fermentation of carbohydrates and their dissimilation via phosphate esters. The most important classes of kinases for phosphorylation are the aldokinases and ketokinases. Hexokinase, one of the most widely studied aldokinases, is responsible for the phosphorylation of glucose, fructose, or mannose, initiating glycolysis. While hexokinase has been found in animals, plants, and bacteria, the similar functioning enzyme glucokinase, which only acts on glucose, has been found in only animals and bacteria. Glucokinase is specific for the conversion of glucose to glucose 6-phosphate and has been reported in a variety of organisms functioning in both aerobic and anaerobic conditions. As verifiable by comparison of K m values for hexokinase and glucokinase, under high glucose concentrations glucokinase has a higher affinity/activity thus is the primary mechanism for glucose assimilation. When galactose concentrations are high, hexokinase is inefficient in carbohydrate phosphorylation, thus a unique enzyme galactokinase is utilized performing a unique phosphorylation of the C1 carbon forming galactose 1-phosphate. This is then followed by an epimerization of galactose 1-phosphate to ­glucose 1-phosphate, which then gets metabolized into glucose phosphates for further conversion into other carbohydrate intermediates. Fructokinase exists in animal and bacterial cells where its activity requires the presence of magnesium and potassium ions. The product of the fructose phosphorylation by fructokinase is fructose 6-phosphate, a central metabolite of carbon flow feeding important pathways such as the lower half of glycolysis and amino sugar synthesis. Another critical group of kinases are referred to as pentokinases because they actively phosphorylate pentoses specifically at the fifth carbon and have been widely studied in a variety of organisms [4–6,16,22,23,26]. Ribokinase is one of the most important pentokinases since the product of the phosphorylation is ribose 5-phosphate, an important molecule in the pentose phosphate pathway that leads to synthesis of purines, pyrimidines, and histidine.

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3.4.3.2  Aldolases for Monosaccharides Aldolases are abundant enzymes utilized throughout the metabolic network for synthesizing and breaking down metabolites for the generation of carbohydrates as well as energy, nucleic acids, amino acids, and other vital metabolic components. There are two types of aldolases (Type I and Type II) in which each performs the same condensation reaction between an aldehyde acceptor and a ketone donor but where each enzyme tends to have specificity toward a unique donor molecule. Type I aldolases are primarily found in plants and high order animals. They act through a mechanism forming a covalently bound intermediate molecule, after the acceptor molecule creates a Schiff base with the donor molecule. In contrast, Type II aldolases use metal cofactors, specifically Zn2+ , as Lewis acids in the enzyme’s activation site to help facilitate the acceptor–donor transfer and thus no true intermediate is formed. While exceptions are known, typically Type II aldolases are only found in microorganisms. Fructose 1,6-diphophate (FDP) aldolase is one of the most important enzymes in the metabolism of monosaccharides since it lies in the center of glycolysis. The aldolase catalyzes the reaction of glyceraldehyde 3-phosphate and dihydroacetone phosphate (DHAP) to form FDP, the central metabolite leading to pyruvate metabolism and the TCA. FDP aldolase is found in both Type I and Type II acting forms, with Type I generally having tetrameric form and Type II as dimers [31]. Type I FDP aldolases have a high degree of overall sequence homology (greater than 50%) in which the active site is highly conserved throughout the evolution of the enzyme [29]. On the other hand, Type II FDP aldolases have a much lower sequence homology yet studies still suggested similar range in substrate specificity as compared with corresponding Type I substrate specificities. Similar DHAP dependent aldolases included the Type II aldolases for fuculose 1-phosphate and rhamnulose 1-phosphate. Both are highly common Type II aldolases found in microorganisms and function using a similar mechanism where DHAP is the donor for the aldol reaction. These aldolases have been used in whole-cell fermentation systems synthesizing carbohydrates for industrial production. It has also been noted that the use of rhamnulose 1-phosphate can form L-rhamnose, a rare metabolite, by rhamnose isomeration in the cell after the initial aldol reaction forms L-rhamnulose from rhamnulose 1-phosphate [11]. DHAP-dependent aldolases are critical to the availability of particular isomers for later metabolic processes where the enzyme’s in vivo availability is regulated by the simple presence of FDP, fuculose 1-phosphate or rhamnulose 1-phosphate in the cell. A more unique and widely studied aldolase is 2-deoxyribose 5-phosphate (DERA) which has been ­characterized in both function and structure [9,15]. It is unique in that there is no other known aldolase that uses an aldehyde as the donor in the aldol reaction, however the DERA enzyme from E. coli has been found capable of accepting a wide range of donor substrates, however the rate of reaction for more exotic substrates is much slower [1]. Sialic acids are generated solely from N-acetylneuraminate aldolase, a Type I aldolase found in both bacteria and animals. The aldolase catalyzes the condensation of its only known acceptable donor, pyruvate, with acetylmannosamine to form sialic acid [17]. This class of acids and their derivatives, found in bacteria and at the C-terminus of mammalian glycoconjugates, play important roles in biochemical recognition within the cell [27]. Polysialic acids are present in bacterial cells and in mammalian cell tissues where they are believed to be involved in cell adhesion and cell–cell communication [18].

3.5  Cell Synthesis of Lipids 3.5.1  Introduction to Lipids and Fatty Acids The biosynthesis of lipids and fatty acids is a complex interaction of multiple pathways beginning with the generation of free palmitate. In the following steps, palmitate undergoes separate elongation and/or unsaturation to yield the array of fatty acid molecules. Lipids in general are classified similarly to carbohydrates as simple or complex, based on the physical properties of the molecule. Lipids play indispensable roles from cell proliferation and cell differentiation to even organ morphogenesis; all are processes that can be considered intimately associated with cell cycle progression [10]. Apart from serving as

3-26

Cellular Metabolism Sucrose 1 HO—CH2

6 HOCH2 HO

5

O

H

4 H

OH 3 H

H 2

O 1

2

H

O

4

5

6 C—OH 4 H 2

OH

Lactose

5

O

H

HO

OH 3 H

1 O

H

H 2

OH

O

2

3 H

5

O

OH

H

H 1

3 H

OH

2

OH

HO

6 HO-CH2 5

H

4

H 4

H

OH

6 HO-CH2

H 1

H

6 HO-CH2 HO

O

5

H

H HO

3

OH

Maltose

6 HO-CH2

O

OH

4

1 OH 3 H

H 2

H

HO

Figure 3.14  Major disaccharides involved in carbohydrate metabolism. Sucrose, consisting of D-glucose and D-fructose monomers, and maltose, consisting of two D-glucose monomers, both having carbon rings linked by C1 α anomer bonds. The D-galactose and D-glucose monomers of lactose have a β linkage.

signaling molecules to regulate developmental processes and defensive mechanisms [12,13], fatty acids and lipids primarily function as energy storage and provide essential components for membrane assembly. Figure 3.14 shows a couple important disaccharides used in energy storage and lipid synthesis. The main mode for new generation of fatty acids occurs in the cytosol starting with the molecule acetyl-CoA and the enzyme acetyl-CoA carboxylase (ACC). 3.5.1.1  Classification of Lipids The heterogeneous group of lipids includes fats, oils, steroids, waxes, and related compounds that all have similar physical properties even though their chemical formulations may vary in structure and composition. Lipids, for the most part, tend to be soluble in nonpolar solvents such as chloroform and ether and are relatively insoluble in water. In many animals they are dietary essential molecules because they are not only a source of energy but also contain important fat-soluble vitamins and other fatty acids found in natural foods. The basic lipid structure classification can be broken into three primary groups. The first group is the simple lipids of fats and waxes, which are generally esters of fatty acids containing various glycerols and alcohols, respectively. The second group is the complex lipids, which are simple lipids containing additional functional groups. For example, the phospholipids are fats in which the molecule includes a phosphoric acid residue sometimes harboring a nitrogen-containing base. Another complex lipid, the glycolipid, carries a carbohydrate group. The third category is the derived lipids. This diverse class of molecules includes steroids, hydrocarbons, hormones, and soluble vitamins among others. 3.5.1.2  Diversity of Lipids in Microorganisms Comparison of lipid composition within a single phylogenetic branch has shown to be similar yet unique to each species, and thus highly useful as marker molecules, especially in chemotaxonomy. The differences in lipid composition are attributed to the variance in growth temperatures, environmental

Biosynthesis of Cellular Building Blocks: The Prerequisites of Life

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nutrient gradients and other factors in the surrounding habitat of each microorganism where the results of which manifest in cellular structures. For example the presence of neutral lipids in membranes, for instance the presence of small amounts of squalene in Archaea and hopanoids in bacteria yet both families contain varying amounts of cartenoids. Additionally, Gram-negative bacteria contain variations within their own outer membrane where each layer differs in composition. For example the E. coli membrane consists of two layers, both of which contain lipoproteins. However, the outer layer is composed of a varying amount of lipopolysaccharides while the inner layer has a composition of phospholipids similar to that of the cytoplasmic membrane. In general, lipids account for about 10% of the cellular building blocks in microorganisms, with roles in energy storage, membrane structure, metabolite transport, and other metabolic processes. The fact that lipids vary in both synthesis route and structure is directly attributable to the different precursor groups used by the various prokaryotes. The basic components of most cellular lipids are

1. Glycerol phosphate, which is formed from DHAP through a reduction reaction. 2. Small hydrocarbon chain molecules from the elongation of acetyl-CoA. The chain molecules are usually 14–18 carbon fatty acids in bacteria and 20 carbon isoprene alcohols in Archaea. In both, the small acyl carrier protein (ACP) is used to activate these molecules via thioesterification. 3. A polar head group, which is attached to the long hydrocarbon chain. The two major head groups are glycerol phosphate and ethanolamine phosphate, while others include serine phosphate and glucose.

3.5.2  Sources of Metabolites for Lipogenesis The availability of acetyl-CoA and NADPH plays a vital role in the biosynthesis of fatty acids. AcetylCoA is generated through carbohydrate oxidation to pyruvate, which is then converted to acetyl-CoA by two different pathways. In eukaryotes, pyruvate is first transported into the mitochondria and converted into citrate, which is excreted. Next, using ATP and CoA moieties, citrate is converted into cytosolic acetyl-CoA by an ATP-citrate lyase, which then can serve as the substrate for lipogenesis. Importantly, citrate catabolism in the mitochondria leads to the formation of acetyl-CoA. However, because the mitochondria membrane has no transport mechanisms to move acetyl-CoA into the cytosol, it is unusable for fatty acid biosynthesis. On the other hand, prokaryotes lack mitochondria and thus the synthesis of acetyl-CoA from pyruvate by pyruvate dehydrogenase can be utilized directly for fatty acid synthesis. The cofactor NADPH acts as the proton donor for the reductions of the 3-ketoacyl and 2,3­u nsaturated acyl derivatives in fatty acid biosynthesis. The metabolites produced in oxidative reactions of the pentose phosphate pathway (see Section 3.4.2.1) are the principal source of the hydrogen used for the reductive synthesis of fatty acids. It can be noted that tissues in animals that specialize in the fatty acid synthesis, i.e., the liver, also possess highly active pentose phosphate pathways. Also of note, for all organisms, is that both pathways are located in the cytosol of the cell thus eliminating any permeability barriers that would complicate NADPH transport from one pathway to the other. NADPH is also derived from other sources including the malic enzymes, or NADP malate dehydrogenase, a TCA cycle enzyme used to convert malate to pyruvate.

3.5.3  Synthesis of Long-Chain Fatty Acids From a chemistry perspective, the main oxidative pathway appears to be the opposite route by which fatty acids are synthesized. However, the synthesis of fatty acids takes place in the cytoplasm only using the cofactor NADPH, whereas oxidation occurs in the cytoplasm for prokaryotes and within the mitochondria of eukaryotes utilizing cofactors FADH+ and NAD+ . The Type II, or dissociated fatty acid synthase, is used for microbial fatty acid synthesis to much the same way that polypeptide synthesis occurs in bacteria. Separate proteins are used to catalyze every one of the individual reactions where each protein is encoded by separate genes. This is a stark contrast to the multifunctional protein complexes used

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Cellular Metabolism

in yeast, animals, and birds in which separate protein domains execute reaction steps, a mode referred to as Type I fatty acid synthesis. Regardless of the type of synthesis, there are similarities in synthesis build-up, initiation, elongation, and radical formation. 3.5.3.1 Entrance to the Fatty Acid Pathway Production of malonyl-CoA is the initial step leading to fatty acid synthesis. In addition to NADPH, lipogenesis requires the cofactors ATP, biotin, manganese ions, and bicarbonate for proper functioning of enzymes. Specifically, the carboxylation reaction performed by ACC, in the presence of ATP, converts acetyl-CoA into malonyl-CoA by using bicarbonate as a source of CO2. ACC in E. coli is a multifunctional protein, encoded by accABCD, consisting of biotin carboxylase, biotin acetyl carrier protein (BCCP), transcarboxylase, and a regulatory site. The activation of ACC requires biotin attachment at a specific lysine residue of the BCCP subunit (accBC) by a biotin carrier protein, the biotin ligase (BirA). The A and D subunits (accAD) can then perform the carboxylation reaction; this can also act as a means of competitive regulation. ACP is then incorporated into the active enzyme complex replacing the CoA. It is important to note that malonyl-CoA is a pivotal metabolite in plant biosynthesis. This is because there are two branches for which carbon flow is then distributed: (1) to the synthesis of fatty acids and (2) to the synthesis of flavonoids, the multifunctional secondary metabolites that confer plant colorations, antimicrobial protection and UV protection. In humans, flavonoids are believed to have high medicinal value as cancer preventive, antipathogens, and antidiabetic. The increasing attention of flavonoids as nutraceuticals has driven many biotechnological endeavors to produce these metabolites from E. coli and S. cerevisiae using these organism’s native malonyl-CoA as the starter unit for flavonoid synthesis [19–21,32,33]. 3.5.3.2  Initiation, Radical Formation, and Liberation Following malonyl-CoA formation, a condensation of malonyl-CoA by malonyl-ACP transacylase occurs as a first step toward fatty acid synthesis (Figure 3.15). The malonyl moiety reacts with a free ACP forming malonyl-ACP. The free CoA byproduct is then recycled back into the metabolic network for later use. Some malonyl-ACP molecules can then be recycled back to acetyl-CoA via acetyl-ACP (through enzymes encoded by fabB and fabF in E. coli), whereas the remaining pool enters into a multistep cycle where two carbon atoms are added each cycle through addition of acetyl groups creating fatty acid polymers. The fatty acid biosynthesis initiation is marked by one of two committed steps: (1) the formation of acetyl-ACP from malonyl-ACP and acetyl-CoA or (2) the formation of acetoacetyl-ACP from malonyl-ACP and acetyl-ACP. These molecules, acetyl-ACP and acetoacetyl-ACP, are sometimes referred to as the primer molecules since the elongation cycle builds the hydrocarbon chain off these molecules. Branched-chain fatty acids are synthesized using a primer molecule formed by either of two methods: in one, malonyl-ACP is joined with an aldehyde generated from intermediates of branchedchain amino acid synthesis. The other method uses an alternative primer molecule formed by a specialized transcylase(s) that reacts a free ACP with a lone acyl-CoA. The final long-chain radical is formed through an elongation of the prosthetic group attached to the ACP by the series of reduction, dehydration, and condensation processes. The four-step cycle of elongation begins with the condensation of the acetyl group on the primer molecule releasing carbon dioxide and ACP to form 3-ketoacyl-CoA. The 3-ketoacyl-ACP synthase isoenzymes are responsible for catalyzing the irreversible reaction to elongate the carbon chain. In E. coli, the enzyme responsible is acetyl-CoA ACP transcylase (fabH). Next the 2-oxo group is reduced to form a D-β-hydroxyl group by 3-ketoacyl-CoA reductase, forming 3-hydroxyacyl-CoA using the cofactor NADPH; in bacteria this is generally performed by 3-oxoacyl-ACP reductase (fabG). The first reduction is followed by a dehydration via 3-hydroxyacyl-CoA dehydratase, or β-hydroxybutyryl ACP dehydratase (fabD) in E. coli, that yields 2-trans-enoyl-CoA. Finally, a second reduction completes the elongation cycle to form another saturated acyl-ACP that has been lengthened by two carbon atoms. Enoyl-ACP reductase (fabF), a

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Biosynthesis of Cellular Building Blocks: The Prerequisites of Life (a)

O CH3

C

ACP

S

CH2

H3C

1

O HOOC

C

O

O

Acetyl -ACP

S

ACP

ACP C S CH2 Acetoacetyl -ACP (primer) C

Malonyl -ACP (b)

O CH2 C [CH2]x CH2

H3C

S

ACP

4

5

ACP-SH + CO2

Malonyl -ACP 2 [H] O H3C

H2O

O

O

CH C [CH2]x CH

S

ACP

H3C

S

ACP

2 [H]

3

2 OH

H3C

C C [CH2]x CH2

O

C C [CH2]x CH2

(c)

S

ACP

10:0 β -Hydroxydecanoyl -ACP 6 6 10:1 ∆3 cis-3-Decanoyl -ACP 7

10:1 ∆2 trans-2-Decanoyl -ACP

7 or 8

16:1 ∆9 Palmitoleyl -ACP 8

18:0 Palmitoyl -ACP

18:1 ∆11 cis-Vaccenoyl -ACP

Figure 3.15  Fatty acid biosynthesis cycle for long chain polysaccharides. (a) Initiation begins with the formation of a primer molecule via a number of different 1, β-ketoacyl-ACP synthetases. (b) The elongation cycle used to add malonyl moieties is conducted by 2, β-ketoacyl-ACP reductase; dehydration by 3, β-hydroxylacyl-ACP dehydrase; reduction by the NADPH-dependent 4, enoyl-ACP reductase; and condensation of the next malonyl moiety by a 5, β-ketoacyl-ACP synthetase. (c) Distribution into different classes of long chain fatty acids is performed by 6, b-hydroxydecanoyl-ACP dehydrase (HDD) catalyzing the dehydration and a 3-cis isomer isomerization; 7, β-ketoacyl-ACP synthase isoenzyme I; β-ketoacyl-ACP synthase isoenzyme II.

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Cellular Metabolism

2-trans-enoyl-CoA reductase, with a reducing cofactor NADPH forms the saturated carbon chain that can then serve as the substrate for the next elongation cycle. For small saturated fatty acid chains (<10 carbons in length), the process of condensation, reduction, and dehydration continues until the chain reaches a length of ten carbons. There are a couple of routes to elongate the fatty acid chain further: β-hydroxydecanoyl-ACP dehydrogenase (HDD) can catalyze either a single dehydration or performed along with an isomerization reaction. The isomerization utilizes only the 2-trans form converting the chain to its 3-cis isomer, thus creating the unsaturated form of the fatty acid. When only undergoing the dehydration reaction, ketoacyl-ACP synthase (KAS) I or its isoenzyme KAS II continues to grow the chain until free palmitate, a 16 carbon polypeptide, is formed. The overall synthesis equation from acetyl-CoA and malonyl-CoA to palimate is

CH2CO.S.CoA + 7 HOOC.CH2CO.S.CoA + 14 NADPH + 14 H + → CH3(CH2)14COOH  + 7 CO2 + 6 H2O + 8 CoA.SH + 14 NADP + 

In the case of isomerization, 2-trans-enoyl-CoA reductase is not capable of performing the reduction reaction, thus forcing the further elongation towards unsaturated fatty acids. For the synthesis of unsaturated fatty acids, the C10-ACP intermediate is used as the precursor for elongation continuing by only KAS I until the chain has reached sixteen carbons (C16:1Δ9 cis) in length. From there, KAS II actively elongates the unsaturated fatty acid to a chain length of 18 carbons (C18:1Δ11 cis). This so-called anaerobic pathway is used for synthesis of unsaturated fatty acids in anaerobic microbes and in many aerobic microbes. In many Gram-positive aerobes, a so-called aerobic pathway is used for the generation of polyunsaturated fatty acids. In this pathway the unsaturation occurs not at 10 carbons but at 16 carbons in length. A fatty acid desaturase, with the help of molecular oxygen, introduces a double bond in the middle of C16 acyl-ACP to form C16:1Δ9 cis (palmitoleic acyl-ACP). 3.5.3.3  Completion of Lipid Synthesis As the acyl chains are formed through the elongation process, the acyl-ACP is transferred into membrane phospholipids by glycerol phosphate acyltransferase (encoded by plsB), which yields the phosphatic acids. Different acyltransferases have preference for different acyl chain lengths and cis-unsaturated or branched-chain acids, thus ensuring an even distribution of different fatty acids in the membrane. The addition of polar head groups follows the insertion of the fatty acid molecule into the membrane by a variety of transfer mechanisms to produce a phospholipid or one of the other major classes. The most prevalent phosphatic acid, diacylglycerol 3-phosphate, is activated by nucleotidyl transfer from CTP by CDP-diglyceride synthase to form a CDP-diacylglycerol. Subsequent reactions by a variety of specific enzymes lead to the array of phospholipids. Diacylglycerol 3-phosphate can also be activated by glycosyl transfer to after a phosphorylation to give the precursor for glycolipids.

3.5.4  Synthesis of Neutral Lipids Neutral lipids are a group of lipophilic compounds (i.e., carotenoids, squalene, C 40 isoprenoids) derived from acetyl-CoA via addition of C5-isoprene units. This altered synthesis route offers a distinct change in the structure and function between neutral lipids and traditionally formed fatty acids. Like most of the long-chain saturated and unsaturated fatty acids making up the bulk of cellular lipids, neutral lipids are also essential structural and functional elements of membranes. These polymers are known to play roles in membrane stability, electron transport, sugar, and oligosaccharide transfer through membranes, and even in light absorption for protection from light or to harvest light energy (specifically in bacteria). Much of the current literature shows the importance of polar lipids, i.e., phospholipids, while comparatively little attention has been paid to the metabolism of neutral lipids. An excellent review of neutral lipids has been assembled by Coppens and Vielemeyer that includes a discussion of new pharmacological targets from pathways generating neutral lipids [7].

Biosynthesis of Cellular Building Blocks: The Prerequisites of Life

3-31

As a first step in their biosynthetic pathway, two acetyl-CoA molecules are condensed to form a single acetoacetyl-CoA. A third molecule of acetyl-CoA is then quickly condensed on the carbon of the second ketone group, creating 3-hydroxy-3-methyl-glutaryl-CoA (HMG-CoA). A reduction of the CoA-activated acid to an alcohol occurs to form mevalonic acid, which in turn is diphosphated to from a single isoprenyl unit, or isopentenyl diphosphate. From here one isoprenyl unit is isomerized, forming a prenyl unit, to which a second isoprenyl unit is condensed onto it, thus creating a chained molecule via a simple polymerization chain reaction. This process can continue to create very large molecules; such is the case for C55 isoprenoids and quinones. The polymerization process is generally characterized by reference to head and tail units of each molecule where the product is stabilized by hydrogen ion elimination. The pyrophosphate leaving group pulls out the oxygen atom during the activation process to leave behind a reactive carbenium cation, referred to as the head. This head can easily add to the tail, or the double bond of the electrophilic ­isopentenyl unit, of an adjacent molecule. The condensation reaction can occur in conformations of head-to-tail (as explained above), head-to-head, or tail-to-tail to form the variety polyprenyl compounds containing two, three, four or more isoprenoid units. Those with more than four units are generally derived by the union of two sesquiterpene (being six units in length) or two diterpene units (having a total of eight units).

References 1. Barbas, C. F., Y. F. Wang, and C. H. Wong. 1990. Deoxyribose-5-phosphate aldolase as a synthetic catalyst. J. Am. Chem. Soc., 112:2013–14. 2. Buckley, R. H. 2002. Primary cellular immunodeficiencies. J. Allergy Clin. Immunol., 109:747–57. 3. Buckley, R. H. 2002. Primary immunodeficiency diseases: dissectors of the immune system. Immunol. Rev., 185:206–19. 4. Cabrera, R., A. Caniuguir, A. L. Ambrosio, V. Guixe, R. C. Garratt, and J. Babul. 2006. Crystallization and preliminary crystallographic analysis of the tetrameric form of phosphofructokinase-2 from Escherichia coli, a member of the ribokinase family. Acta Crystallograph Sect. F Struct. Biol. Cryst. Commun., 62:935–37. 5. Cao, P., Y. Gong, L. Tang, Y. C. Leung, and T. Jiang. 2006. Crystal structure of human pyridoxal kinase. J. Struct. Biol., 154:327–32. 6. Chuvikovsky, D. V., R. S. Esipov, Y. S. Skoblov, L. A. Chupova, T. I. Muravyova, A. I. Miroshnikov, S. Lapinjoki, and I. A. Mikhailopulo. 2006. Ribokinase from E. coli: expression, purification, and substrate specificity. Bioorg. Med. Chem., 14:6327–32. 7. Coppens, I. and O. Vielemeyer. 2005. Insights into unique physiological features of neutral lipids in Apicomplexa: from storage to potential mediation in parasite metabolic activities. Int. J. Parasitol., 35:597–615. 8. Demuynck, C., F. Fisson, I. Bennanibaiti, H. Samaki, and J. C. Mani. 1990. Immunoaffinity purification of transketolases from yeast and spinach leaves. Agri. Biol. Chem., 54:3073–78. 9. DeSantis, G., J. Liu, D. P. Clark, A. Heine, I. A. Wilson, and C. H. Wong. 2003. Structure-based mutagenesis approaches toward expanding the substrate specificity of D-2-deoxyribose-5-phosphate aldolase. Bioorg. Med. Chem., 11:43–52. 10. Donnelly, P. M., D. Bonetta, H. Tsukaya, R. E. Dengler, and N. G. Dengler. 1999. Cell cycling and cell enlargement in developing leaves of Arabidopsis. Dev. Biol., 215:407–19. 11. Drueckhammer, D. G., J. R. Durrwachter, R. L. Pederson, D. C. Crans, L. Daniels, and C.-H. Wong. 1989. Reversible and in situ formation of organic arsenates and vanadates as organic phosphate mimics in enzymatic reactions: mechanistic investigation of aldol reactions and synthetic applications. J. Org. Chem., 54:70–77. 12. Farmer, E. E. 1994. Fatty acid signalling in plants and their associated microorganisms. Plant Mol. Biol., 26:1423–37.

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13. Farmer, E. E., H. Weber, and S. Vollenweider. 1998. Fatty acid signaling in Arabidopsis. Planta, 206:167–74. 14. Fraenkel, D. G. 1986. Mutants in glucose metabolism. Annu. Rev. Biochem., 55:317–37. 15. Horinouchi, N., J. Ogawa, T. Sakai, T. Kawano, S. Matsumoto, M. Sasaki, Y. Mikami, and S. Shimizu. 2003. Construction of deoxyriboaldolase-overexpressing Escherichia coli and its application to 2-deoxyribose 5-phosphate synthesis from glucose and acetaldehyde for 2´-deoxyribonucleoside production. Appl. Environ. Microbiol., 69:3791–97. 16. Jeffries, T. W. and Y. S. Jin. 2004. Metabolic engineering for improved fermentation of pentoses by yeasts. Appl. Microbiol. Biotechnol., 63:495–509. 17. Kim, M.-J., J. Hennen, H. M. Sweers, and C.-H. Wong. 1988. Enzymes in carbohydrate ­synthesis: N-acetylneuraminic acid aldolase catalyzed reactions and preparation of N-acetyl-2-deoxy-Dneuraminic acid derivatives. J. Am. Chem. Soc., 110:6481–86. 18. Lasky, L. A. 1992. Selectins: interpreters of cell-specific carbohydrate information during inflammation. Science, 258:964–69. 19. Leonard, E., J. Chemler, K. H. Lim, and M. A. Koffas. 2006. Expression of a soluble flavone synthase allows the biosynthesis of phytoestrogen derivatives in Escherichia coli. Appl. Microbiol. Biotechnol., 70:85–91. 20. Leonard, E., Y. Yan, and M. A. Koffas. 2006. Functional expression of a P450 flavonoid hydroxylase for the biosynthesis of plant-specific hydroxylated flavonols in Escherichia coli. Metab. Eng., 8:172–81. 21. Leonard, E., Y. Yan, K. H. Lim, and M. A. Koffas. 2005. Investigation of two distinct flavone synthases for plant-specific flavone biosynthesis in Saccharomyces cerevisiae. Appl. Environ. Microbiol., 71:8241–48. 22. Long, M. C., V. Escuyer, and W. B. Parker. 2003. Identification and characterization of a unique adenosine kinase from Mycobacterium tuberculosis. J. Bacteriol., 185:6548–55. 23. Maugeri, D. A., J. J. Cazzulo, R. J. Burchmore, M. P. Barrett, and P. O. Ogbunude. 2003. Pentose phosphate metabolism in Leishmania mexicana. Mol. Biochem. Parasitol., 130:117–25. 24. McCarthy, G. 2004. Medical diagnosis, management and treatment of Lesch Nyhan disease. Nucleosides Nucleotides Nucleic Acids, 23:1147–52. 25. Nyhan, W. L. 2005. Disorders of purine and pyrimidine metabolism. Mol. Genet. Metab., 86:25–33. 26. Parducci, R. E., R. Cabrera, M. Baez, and V. Guixe. 2006. Evidence for a catalytic Mg2+ ion and effect of phosphate on the activity of Escherichia coli phosphofructokinase-2: regulatory properties of a ribokinase family member. Biochemistry, 45:9291–99. 27. Paulson, J. C. 1989. Glycoproteins: what are the sugar chains for? Trends Biochem. Sci., 14:272–76. 28. Suami, T. 1979. Modifications of aminocyclitol antibiotics. Jpn. J. Antibiot., 32 Suppl:S91–102. 29. Sygusch, J., D. Beaudry, and M. Allaire. 1987. Molecular architecture of rabbit skeletal muscle aldolase at 2.7-A resolution. Proc. Natl. Acad. Sci. USA, 84:7846–50. 30. Tsukada, Y., W. K. D. Bischof, N. Hibi, H. Hirai, E. Hurwitz, and M. Sela. 1982. Effect of a conjugate of daunomycin and antibodies to rat alpha-fetoprotein on the growth of alpha-fetoprotein­producing tumor-cells. Proc. Natl. Acad. Sci. USA, 79:621–25. 31. Vonderosten, C. H., A. J. Sinskey, C. F. Barbas, R. L. Pederson, Y. F. Wang, and C. H. Wong. 1989. Use of a recombinant bacterial fructose-1,6-diphosphate aldolase in aldol reactions—preparative syntheses of 1-deoxynojirimycin, 1-deoxymannojirimycin, 1,4-dideoxy-1,4-Iimino-D-arabinitol, and fagomine. J. Am. Chem. Soc., 111:3924–27. 32. Yan, Y., J. Chemler, L. Huang, S. Martens, and M. A. Koffas. 2005. Metabolic engineering of anthocyanin biosynthesis in Escherichia coli. Appl. Environ. Microbiol., 71:3617–23. 33. Yan, Y., A. Kohli, and M. A. Koffas. 2005. Biosynthesis of natural flavanones in Saccharomyces ­cerevisiae. Appl. Environ. Microbiol., 71:5610–13. 34. Zhao, J., T. Baba, H. Mori, and K. Shimizu. 2004. Global metabolic response of Escherichia coli to gnd or zwf gene-knockout, based on 13C-labeling experiments and the measurement of enzyme activities. Appl. Microbiol. Biotechnol., 64:91–8.

4 Polymerization of Building Blocks to Macromolecules: Polyhydroxyalkanoates as an Example Si Jae Park LG Chem Ltd.

Soon Ho Hong University of Ulsan

Sang Yup Lee Korea Advanced Institute of Science and Technology

4.1 4.2 4.3

Introduction ���������������������������������������������������������������������������������������4-1 PHAs ���������������������������������������������������������������������������������������������������4-2 PHA Synthases �����������������������������������������������������������������������������������4-2

4.4

 etabolic Engineering of Microorganisms for PHA M Production ����������������������������������������������������������������������������������������� 4-5





Evolution of PHA Synthases

SCL-PHA  •  MCL-PHA and SCL-MCL-PHA

4.5 Conclusion ����������������������������������������������������������������������������������������4-14 Acknowledgments ��������������������������������������������������������������������������������������4-15 References ����������������������������������������������������������������������������������������������������4-15

4.1  Introduction Microorganisms synthesize many building blocks that are mainly used for the synthesis of macromolecules such as proteins, DNA, RNA, lipids, and so on. Because these macromolecules are indispensable for living organism, the metabolism is optimized to provide enough precursors to synthesize them. A polymer is made by covalently linking together simple small molecules called monomers. A general term to describe the process that leads to the formation of a polymer is polymerization. It should be noted that there are many ways to polymerize monomers to synthesize various macromolecules. Biopolymers are amongst the most sophisticated and complex polymers on earth, and it is important to understand how monomers or building blocks can assemble covalently into life-enabling polymers. There are many different biopolymers that have been studied in detail with respect to their biosynthetic mechanisms, enzymes, and precursors involved in, and metabolic engineering strategies. For detailed information on various biopolymers, it is recommended for readers to refer to the excellent book on biopolymers recently published [1]. Since this book is about metabolic engineering and only limited space is available, we focus on only one family of biopolymers for detailed description. Polyhydroxyalkanoates (PHAs) are an interesting family of carbon and energy storage polymers, which are formed by polymerization of monomers derived from central carbon metabolism [2]. There have been many advances in our understanding of the metabolism of PHA biosynthesis and the molecular characteristics of the genes and enzymes involved. A number of interesting metabolic engineering studies have been performed toward enhanced production of PHAs, production of novel PHAs, and utilization of inexpensive carbon sources. We will start with what PHAs are and how they are 4-1

4-2

Cellular Metabolism

synthesized. Following a description of the metabolism and molecular characteristics of the genes and enzymes involved, we will review the metabolic engineering examples. Even though the polymerization processes are different for other biopolymers, the general strategies described in this chapter should be valid for metabolic engineering toward enhanced production of biopolymers and for the production of novel and/or tailor-made biopolymers.

4.2  PHAs Among various biopolymers produced by microorganisms, PHAs have attracted much attention for commercial application due to mechanical properties that are similar to general-purpose polymers and their complete biodegradability. The monomer constituents of natural PHAs identified to date are more than 150 hydroxyalkanoic acids [3]. (R)-3-hydroxyalkanoic acids are most frequently used as monomers of PHAs. PHAs show a broad range of polymer characteristics from thermoplastics to rubber like polymers depending on the types and amounts of monomer constituents [4]. Thermoplastic short chain length (SCL) PHAs are composed of monomers with three to five carbon atoms, and rubberlike medium chain length (MCL) PHAs are composed of monomers with six to 14 carbon atoms [4]. Recently, PHA copolymers containing both SCL and MCL monomers (SCL-MCL-PHA) were identified in several bacteria. Material properties of these copolymers are dependent on the amount of SCL or MCL monomers. For example, incorporation of small amounts of MCL monomer units into poly(3hydroxybutyrate) [P(3HB)] backbone polymers makes the properties of the PHA material similar to low density polyethylene (LDPE), which opens new opportunities for commercial applications of PHAs [5]. PHA synthesis in microorganism can be divided into two major processes: (R)-hydroxyacyl-CoAs (RHA-CoAs) are generated from various metabolic pathways and then RHA-CoAs are polymerized into PHA. In a broad sense, all metabolic pathways of microorganism are involved in these processes to provide RHA-CoAs that are finally used up for PHA synthesis. However, when we consider enzymes directly involved in PHA biosynthesis, enzymes can fall into three classes according to their function. The first group of enzymes are PHA synthases that have a key role in PHA biosynthesis, the polymerization of RHA-CoAs (Figure 4.1; Table 4.1). Wide substrate specificity of PHA synthases results in the corporation of the variety of monomer constituents in PHAs. The second group consists of the enzymes that generate RHA-CoAs from the intermediates of metabolism, such as β-ketothiolase, 3-ketoacyl -CoA reductase, and enoyl-CoA hydratase (Table 4.2). Finally, enzymes involved in the regulation of PHA synthesis at the transcription and translation level and in stabilization of PHA granules in bacteria, such as PhaR, PhaF, and phasin (PhaI and PhaP) correspond to the third group.

4.3  PHA Synthases To synthesize PHA in the cytoplasm of microorganisms, the combined action of RHA-CoA generating enzymes and RHA-CoA polymerizing PHA synthase is very critical. PHA synthase carry out the key role in PHA synthesis, the polymerization of RHA-CoAs into PHA granules. To date, over 60 PHA synthases have been obtained and characterized at a molecular level [6]. PHA synthases can be classified into four major groups depending on the substrate specificity, enzyme subunits consisting of PHA synthase and genetic organization type of PHA synthesis operon (Figure 4.1) [6]. Class I PHA synthases have high substrate specificity toward RHA-CoAs having three to five carbon atoms and consist of only one type of subunit, PhaC with molecular weight of around 60–73 kDa. Ralstonia eutropha and Alcaligenes latus PHA synthases belong to Class I group [7,8]. In R. eutropha, PHA synthase gene (phaC) constitutes operon (phaCAB) with phaA and phaB genes encoding β-ketothiolase and NADPH-dependent acetoacetyl-CoA reductase, respectively. A different genetic order of phaC, phaA, and phaB in the PHA operon is often found in some bacteria, but the phaC gene is often found to be colinearized with phaA and phaB genes. Class II PHA synthases are also composed of one type of subunit, PhaC with molecular weight around 60–65 kDa, but have higher specificity to RHA-CoAs having six to 12 carbon atoms. Class II PHA

phaC (1767 bp)

phaQ (441 bp)

phaA (1185 bp)

PhaC (40 kDa)

phaB (744 bp)

PhaR (22 kDa)

phaB (738 bp)

phaB (741 bp)

phaD (615 bp)

phaP (363 bp)

phaC (1089 bp)

ORF4 (462 bp)

phaC2 (1680 bp)

E. coli σ70 consensus promoter

phaR (609 bp)

phaE (1074 bp)

Figure 4.1  Genetic organization of PHA synthases and their subunits.

phaP (513 bp)

Class IV: Bacillus megaterium

phaC (1068 bp)

PhaC (40 kDa)

phaZ (849 bp)

PhaE (40 kDa)

phaA (1179 bp) PhaC (60–65 kDa)

PhaC (60–73 kDa)

σ70- dependent promoter

Class III: Allochromatium vinosum

phaC1 (1677 bp)

Class II: Pseudomonas aeruginosa

E. coli σ70 consensus promoter

Class I: Ralstonia eutropha

ORF7

Polymerization of Building Blocks to Macromolecules 4-3

4-4

Cellular Metabolism

synthases are preferentially found in Pseudomonas sp., for example Pseudonomas aeruginosa [9] and P. putida. Two types of PHA synthase gene (phaC1 and phaC2) constitute the operon, in which phaC1 and phaC2 genes are separated by phaZ gene encoding intracellular PHA depolymerase. Two PHA synthases show high amino acid sequence homology and have similar substrate specificity. Downstream of the phaC1, phaZ, phaC2 operon, genes encoding regulatory proteins such as PhaD, PhaI, and PhaF are colocalized. Class III PHA synthases are often found in cyanobacteria and are firstly cloned and characterized from Allochromatium vinosum [10]. They consist of two different types of subunits, PhaC and PhaE subunits of molecular weight of ca. 40 kDa. PhaC has an amino acid sequence homology of 21–28% with Class I and II PHA synthases but PhaE shows no sequence similarity to PHA synthases. In A. vinosum, the phaCE genes and phaAB genes are located in opposite direction. These PHA synthases prefer to use RHA-CoAs with three to five carbon atoms, however, they were recently found to have rather broad substrate specificity toward RHA-CoAs with three to ten carbon atoms. Class IV PHA synthases exist in Bacillus sp. and are composed of two different types of subunits, PhaR and PhaC [11]. Molecular weights of PhaR and PhaC are 20 kDa and 40 kDa, respectively. The phaP, phaQ, phaR, phaB, phaC genes are colocalized in the chromosome and phaPQ genes and phaRBC genes are in opposite direction. These PHA synthases preferentially use RHA-CoAs with three to five carbon atoms. Substrate specificity of PHA synthases regarding chain length of RHA-CoAs is one of the key factors in classification of PHA synthases groups. Recently, PHA synthases having rather broad substrate specificity were found in some bacteria including Thiocapsa pfennigii [12], Pseudomonas sp. 61-3 [13], and Aeromonas caviae [14]. T. pfennigii has PHA synthases belonging to Class III with substrate specificity to RHA-CoAs comprising three to 12 carbon atoms. PHA synthases found in Pseudomonas sp. 61-3 belong to Class II, however, PhaC1 and PhaC2 from Pseudomonas sp. 61-3 are found to polymerize PHAs consisting of 3-hydroxybutyrate (3HB) and MCL-RHAs. A. caviae PHA synthase showing high similarity to Class I PHA synthase (ca. 45%) synthesize PHAs consisting of 3HB and 3-hydroxyhexanoate (3HHx). Further studies revealed that A. caviae PHA synthase could even accept 3-hydroxyhepanoylCoA (3HHp-CoA) [15]. Furthermore, a typical Class I PHA synthase, R. eutropha PHA synthase, was found to produce a copolyester consisting of 3HB and MCL-RHAs when metabolic pathways of host strains were well optimized to supply MCL-RHA-CoAs [16,17]. All these findings lead us to conclude that substrate specificity for PHA synthases is very versatile and broad.

4.3.1 Evolution of PHA Synthases Biosynthesis of PHA consists of two major steps, monomer substrate supply and polymerization of monomer units by PHA synthase. PHA productivity is a function of the activities of enzymes supplying monomers and polymerization of the monomers. How to efficiently supply monomers is more critical because generally polymerization of PHA is executed only in conditions that substrate specificity and activity of PHA synthase have been well optimized to supplied monomers. In particular, PHA synthase is a key factor in determining PHA productivity as well as polymer material properties including molecular weight, polydispersity, and composition and distribution levels of monomer in copolymer. Therefore, PHA synthase has always been a hot issue in the PHA research field. Studies on the amino acid sequence of PHA synthases obtained from various microorganisms and especially on the polymerization mechanism of PHA synthases from R. eutropha and A. vinosum have provided some clues to understand important amino acid residues responsible for polymerization. As protein engineering studies are generally carried out on the basis of tertiary structure revealed by X-ray crystallography when important amino acid residues are changed through site directed mutagenesis, the lack of tertiary structure information for polymerase hampers any in depth protein engineering studies of PHA synthases. Even though the exact tertiary structure of PHA synthases is not presently available, bioinformatic tools provide a basic idea to assume the structure of polymerase, leading to the design of experimental schemes to find important amino acid residues in PHA biosynthesis. Since the first report about the nucleotide sequence of

Polymerization of Building Blocks to Macromolecules

4-5

R. eutropha PHA synthase, it was found that all PHA synthases contain a lipase box (G-X-[S/C]-X-G) in which the essential active site serine of lipase is substituted by cysteine [7]. Furthermore, an amino acid sequence homology search by BLAST using A. vinosum PhaC as a query supported that cysteine is well aligned with serine in bacterial lipases from P. cepacia, Pseudomonas sp. KW1-56, and P. luteola and also revealed that a span of 45 residues (131–175) including the lipase box of A. vinosum PhaC shows 42% sequence identity to P. cepacia lipase [18]. P. cepacia lipase has been characterized at a crystal structure level, therefore, this structural information is helpful in investigating PHA synthase because PhaC is likely to contain similar structures in this region. Using the P. cepacia lipase tertiary structure, the Sinskey and Stubbe group recently reported the molecular mechanism of polymerization of A. vinosum PHA synthase. Mutagenesis studies generating mutation in A. visnosum PHA synthase C149, H331, H303, D302, and C130 residues revealed that H331 is the general base catalyst that activates the nucleophile, C149, for covalent catalysis [18]. Studies also corrected previously proposed role of C130 in polymerization by showing that C130 is not truly involved in catalysis. Authors suggest D302 functions as a general base catalyst in activation for 3HB-CoA for nucleophilic attack on the covalently linked thiol ester intermediate and affect the chain elongation during polymerization [18]. Due to the lack of structure information of PHA synthase, screening and characterization of newly developed PHA synthase after random mutagenesis have been carried out to investigate the molecular characteristics of polymerase. Mutant PHA synthases from R. eutropha, A. caviae, Pseudomonas sp. 61-3, and P. putida have been generated by random mutagenesis through error-prone PCR and employing a mutator strain [19–31]. Mutant PHA synthases were screened by selecting cell colonies showing enhanced biosynthesis level of P(3HB), the most characterized PHA member. Once PHA synthases showing dramatic changes in P(3HB) accumulation level were screened, site directed saturation mutagenesis of target amino acid were further executed to identify other mutations conferring similar enhanced characteristics. Representative results of in vitro mutagenesis conferring significant effects in biosynthesis of PHA in various polymerases are summarized in Table 4.1.

4.4 Metabolic Engineering of Microorganisms for PHA Production 4.4.1  SCL-PHA Substrates of PHA synthases, RHA-CoAs are generated by activation of hydroxyfatty acid using CoA transferase and CoA synthetase and by conversion of inherent intermediates of host metabolism. The first methods are typically used for the investigation of newly found enzymes, especially PHA synthases obtained from novel PHA producing bacteria, for their activities in PHA biosynthesis and for biosynthesis of PHA consisting of novel monomer units. Usually, hydroxyfatty acid is hard to use for the PHA biosynthesis in large amounts because it is expensive and toxic to the cells. Therefore, it is employed to the small-scale production of novel PHA to confirm the feasibility of process. The recombinant microorganisms expressing CoA transferase or CoA synthetase responsible for the activation of corresponding hydroxyfatty acid have successfully synthesized PHAs containing SCL-monomers such as 3HB, 3-hydroxypropionate (3HP), 4-hydroxybutyrate (4HB), and 3-hydroxyvalerate (3HV). Coexpression of the Clostridium kluyveri 4-hydroxybutyryl-CoA:CoA transferase gene (cat2) and the R. eutropha PHA synthase gene in E. coli lead to the production of P(4HB) homopolymer from glucose and 4-hydroxybutyric acid. In the absence of glucose, P(3HB-co-4HB) containing 72 mol% of 3HB was accumulated, which indicates that E. coli seems to operate hither-to-unknown pathways to convert 4HB to 3HB, that is suppressed in the presence of glucose [32]. 4HB could be activated into 4HB-CoA by simultaneous expression of the Clostridium acetobutylicum butyrate kinase (buk) and phosphotransbutyrylase (ptb) in E. coli [33]. Growing E. coli strain expressing the phaC, ptb, and buk genes in the medium supplemented with 3-hydroxybutyric acid, 4-hydroxybutyric acid, and 4-hydroxyvaleric acid resulted in the formation of homopolymers of 3HB, 4HB, and 4HV, respectively.

Pseudomonas Gpo1 PhaC1 and Pseudomonsa aeruginosa PhaC1

Several point mutation in α/β hydrolase region/Site specific mutation

E130D Q418K S325C Q481M

Pseudomonas sp.61-3 PhaC1

Ralstonia eutropha PhaC

Aeromonas caviae PhaC

F518I F362I, F518I D459V, A513C D171G N149S F420S G4D E267K Y445F L446K

Mutation

Aeromonas punctada PhaC

Enzymes

Characteristics

Increase of PHA content Increase of 3HHx fraction in copolymer Increase of specific activity of PHA synthase/decrease of molecular weight of PHA Increase of molecular weight of PHA/increase of expression level of PhaC Decrease of specific activity of PHA synthase Decrease of specific activity of PHA synthase/conferring substrate specificity toward 3HO to PhaC Decrease of specific activity of PHA synthase Increase of PHA synthesis activity/increase of 3HB fraction in copolymer Increase of molecular weight of PHA Increase of 3HB fraction in copolymer Increase of 3HB fraction in copolymer Change of substrate specificity of PhaC1 to use 3HB as a monomer

Increase of specific activity of PHA synthase Increase of molecular weight of PHA

Table 4.1  Representative Mutagenesis to Change Characteristics of PHA Synthase

Site specific or semi random mutagenesis in specific region

Error-prone PCR

Error-prone PCR/ saturated mutagenesis

Error-prone PCR

Mutator strain

Methods

[30,31]

[19,22,23,25]

[20,21,27,28]

[26]

[29]

Reference

4-6 Cellular Metabolism

Polymerization of Building Blocks to Macromolecules

4-7

Pathways for the synthesis of copolymer of 3HP and 3HB were recently developed in recombinant E. coli. PHA copolymer containing 3HP and 3HB was found to have reduced crystallinity, compared with P(3HB) homopolymer. Salmonella enterica serovar Typhimurium propionyl-CoA synthetase (PrpE) was found to have activity to convert 3-hydroxypropionic acid to 3HP-CoA in recombinant E. coli, the substrate of R. eutropha PHA synthase [34]. When 3-hydroxypropionic acid was present in the medium, recombinant E. coli strain expressing the prpE gene and P(3HB) biosynthesis gene could produce copolymer of 3HP and 3HB. P(3HB), the most ubiquitous member of PHAs, is synthesized from acetyl-CoA by sequential reactions catalyzed by three enzymes: β-ketothiolase (PhaA) condensates two acetyl-CoA molecules to acetoacetylCoA, which is reduced to 3HB-CoA by acetoacetyl-CoA reductase (PhaB) using NADPH as a cofactor, and finally 3HB-CoA is incorporated into the growing chain of P(3HB). Because acetyl-CoA and NADPH are used for the synthesis of P(3HB), the metabolic engineering strategy has been focused on modifying the inherent metabolic pathways of E. coli in order to make acetyl-CoA and reducing power more available for P(3HB) biosynthesis pathway. When glucose is used as a carbon source, acetyl-CoA and NADPH can be efficiently provided from the glycolysis and pentose phosphate (PP) pathways. As a strategy to increase cellular NADPH level, overexpression of the zwf and gnd genes, which encode glucose-6-phosphate dehydrogenase and 6-posphogluconate dehydrogenase, respectively, was examined for P(3HB) biosynthesis [35]. When these enzymes were amplified, NADPH/NADP + ratio increased by six times resulting in the increase of the P(3HB) content from 23 to 41%. However, a closer evaluation suggested that the increase of P(3HB) content was not due to enhanced P(3HB) biosynthesis but rather due to decreased cell concentration. Therefore, it is important to make cellular status suitable for P(3HB) biosynthesis without reduction of cell growth (metabolic activity) to achieve high concentration of P(3HB). Various metabolic and fermentation strategies including host strain selection, use of plasmids of different copy numbers, filamentation suppression, use of different PHA biosynthesis genes, and plasmid stabilization have been employed to achieve enhanced PHA biosynthesis in recombinant E. coli [36–38]. These strategies have successfully developed superior strains having high PHA biosynthesis activity, but they could not overcome the root weak point of trial-and-error procedures. Analysis of whole metabolic status of E. coli during P(3HB) biosynthesis was carried out in systems level using 2-dimensional gel electrophoresis (2DE) and metabolic flux analysis (MFA) based on in silico E. coli metabolic network model [39–41]. Proteome analyses of soluble proteins of recombinant E. coli during P(3HB) biosynthesis found proteome patterns such as high level expression of general heat shock proteins including GroES, GroEL, DnaK, and IbpAB [39]. Also, the importance of acetyl-CoA and NADPH availability for P(3HB) production was confirmed by proteome analysis. It was found that the spot intensities of fructose-bisphosphate aldolase (FbaA) and triosephosphate isomerase (TpiA) in glycolytic pathway were increased during P(3HB) accumulation by recombinant E. coli XL1-Blue harboring plasmid pJC4 containing the A. latus PHA biosynthesis genes. The reason for the increased expression of FbaA and TpiA might be due to that E. coli modified its metabolic fluxes to increase the glyceraldehyde-3-phosphate pool, which is eventually used for P(3HB) synthesis. Based on the results of proteome analysis, we examined the effects of the amplification of triosephosphate isomerase (TpiA) and fructose-bisphosphate aldolase (FbaA) on the P(3HB) biosynthesis in recombinant E. coli W3110 harboring the plasmid containing the Ralstonia eutropha PHA biosynthesis genes (Unpublished results, Park SJ and Lee SY). Amplification of TpiA and FbaA significantly increased the P(3HB) content and concentration. In the case of  TpiA amplification, P(3HB) concentration was increased by seven fold compared with control strain (from 0.89 to 5.98 g/L). These results clearly showed that the amplification of glycolytic pathway enzymes can be a good metabolic engineering strategy for the efficient production of P(3HB) by allowing increased glycolytic pathway flux to make more acetylCoA available for P(3HB) biosynthesis. Proteome analysis also reported that the up-regulation of 2-keto-3-deoxy-6-phosphogluconate aldolase (Eda) catalyzing the final step of Entner–Doudoroff (ED) pathway during P(3HB) biosynthesis [39]. Even though the ED pathway has generally been considered to be inactive under normal growth

4HB

Salmonella

C. acetobutylicum

PrpE

Ptb, Buk

b

Engineered A. caviae PhaJ. Engineered E. coli FabH.

3HB, 4HB 3HP

C. kluyveri

Cat2

a

3HB 3HHx, 3HO, 3HD 3HB, 3HV 3HB, 3HV, 3HHx 3HB, 3HV 3HHx, 3HO, 3HD

E. coli E. coli R. eutropha R. eutropha E. coli E. coli

FabH, FabD eFabHb PhaA PhaB Sdm, YgfG TesA Activation of hydroxyfatty acid to HA-CoA

3HHx, 3HO, 3HD 3HHx, 3HO, 3HD

P. aeruginosa

P. putida

RhlG

Glycolysis Glycolysis TCA cycle Fatty acid biosynthesis Fatty acid β-oxidation

Fatty acid β-oxidation Fatty acid biosynthesis

Fatty acid β-oxidation

Fatty acid β-oxidation Fatty acid β-oxidation

3HHx, 3HO, 3HD 3HHx, 3HO, 3HD

E. coli

E. coli

PaaG, PaaF, YdbU

FabG

PhaG

Fatty acid β-oxidation Fatty acid β-oxidation

3HHx, 3HO, 3HD 3HHx, 3HO, 3HD

E. coli

E. coli

Fatty acid β-oxidation

MaoC

3HHx, 3HO, 3HD

P. aeruginosa

PhaJ1, PhaJ2, PhaJ3, PhaJ4

Fatty acid β-oxidation

Fatty acid β-oxidation

Metabolism

YfcX

3HB, 3HHx 3HB, 3HHx, 3HO, 3HD

A. caviae

ePhaJa

Use of inherent metabolic intermediates

Monomer Composition of PHA

Aeromonads

Representative Sources

PhaJ

Enzymes

Table 4.2  Enzymes for PHA Biosynthesis Used in Recombinant E. coli and Salmonella

[33]

[34]

[32]

[64] [65] [7] [7] [46] [75]

[57–60]

[55]

[54–56]

[70]

[69]

[61]

[49,50]

[52]

[48,51]

Reference

4-8 Cellular Metabolism

Polymerization of Building Blocks to Macromolecules

4-9

condition when glucose is used as a carbon source, it seems to become active to satisfy increased demand for NADPH and acetyl-CoA during P(3HB) production. Activation of Eda during P(3HB) biosynthesis was also supported by MFA and comparison of P(3HB) synthesis activity in wild and eda mutant E. coli strain [40]. In silico E. coli metabolic network was constructed with 154 reversible and 156 irreversible reactions and 295 metabolites and metabolic flux distributions in wild type E. coli and recombinant E. coli producing P(3HB) were compared using this model. MFA results showed that ED pathway flux increased significantly under P(3HB) accumulating condition. These MFA and proteome analysis results lead us to compare the abilities of P(3HB) synthesis in E. coli KS272 and its eda mutant. The P(3HB) concentration and the P(3HB) content were lower in eda mutant E. coli strain than those obtained in a wild strain. Because it has been suggested that different E. coli strains have different P(3HB) biosynthesis activity, therefore, it could be concluded that at least in E. coli KS272 strain, the ED pathway plays an important role in P(3HB) biosynthesis. Analysis of P(3HB) granule-associated proteome in recombinant E. coli showed some different aspects of protein expression pattern than in recombinant E. coli producing P(3HB) [41]. Because E. coli does not naturally synthesize P(3HB), there is no system protecting intracellular biomolecules such as DNA, RNA, and proteins from direct contact with hydrophobic P(3HB) granules. Bacteria naturally accumulating PHA have amphiphilic phasin proteins to cover the surface of P(3HB) granules, thus stabilize intracellular biomolecules. Soluble and insoluble proteome analysis of P(3HB) producing recombinant E. coli clearly showed the negative effect of accumulated P(3HB) to intracellular proteins, for example, reduced synthesis of EF-Tu that is critically involved in protein synthesis [39]. Analysis based on soluble proteome lead us to that total expression level of Ef-Tu is reduced due to the unfavorable condition generated by P(3HB) biosynthesis [39]. However, insoluble proteome analysis of revealed that EF-Tu synthesis level is not reduced, but most of the EF-Tu synthesized is associated with P(3HB) granules [41]. New finding was provided by analysis of insoluble proteome that IbpAB, small heat shock proteins that are associated with inclusion body proteins when external stresses such as heat shock are put into E. coli strains, are associated with P(3HB) granules and function to stabilize intracellular biomolecules as phasin proteins do [41]. It has been reported that phasin expression in E. coli does not have a critical effect in enhancing P(3HB) biosynthesis, but the number of P(3HB) granules are increased with reduced diameters of granules. Proteome analysis of P(3HB) granule associated proteome of recombinant E. coli allowed us to find the presence of IbpAB on the surface of P(3HB) granules and to suggest the roles of IbpAB during P(3HB) biosynthesis, induction of P(3HB) accumulation with increased number of granules having decreased diameter [41]. MFA is a useful tool to evaluate cellular metabolic status and to design engineering strategies to increase the product yield under a given condition. Metabolic and fermentation strategies which have been employed to increase P(3HB) biosynthesis in recombinant E. coli such as oxygen limited fermentation and increased availability of acetyl-CoA and NADPH were proved to be right by MFA [42,43]. It was found that under the oxygen limiting condition, additional carbon flux through pyruvate formate lyase increased without any change of the pyruvate dehydrogenase flux, which is used under normal growth condition, resulting in the accumulation of twice more acetyl-CoA, which was subsequently used for P(3HB) biosynthesis. In this section, the importance of acetyl-CoA and NADPH was mainly discussed. However, P(3HB) biosynthesis is not simply controlled by only one factor, but is affected by many central metabolic pathway fluxes. To design host strain for high P(3HB) production activity, it will be important to optimize the flux distributions in such ways that the flux toward P(3HB) is maximized while the fluxes towards other by-products are minimized. Also, fermentation strategies should be carefully optimized because too early accumulation of P(3HB) results in low final cell concentration (due to the growth inhibition) while too late accumulation of P(3HB) results in low P(3HB) content, both of which result in low P(3HB) productivity. Metabolic engineering strategies for the production PHAs containing other RHA monomers besides 3HB have been developed using inherent metabolic intermediates in recombinant E. coli (Figure 4.2).

Citrate

Lactate

SucD

PHA synthase PHA

4HB-CoA

Succinic acid semialdehyde 4HbD 4HB

Glutamate

3HP-CoA

GabT

GadA

PHA

GABA

PHA synthase

Acrylyl-CoA

Lactyl-CoA

PHA

PHA synthase

FabG Acyl-CoA

FadD

Fatty acids

FabG

Enoyl-CoA

PHA

PHA synthase

(R)-3-Hydroxyacyl-CoA

?

YfcX, YdbU, (S)-3-Hydroxyacyl-CoA PaaG, PaaF MaoC

FadA FadE Fatty acid 3-Ketoacyl-CoA β-oxidation pathway FadB FadB

Thioesterase (TesB)

(R)-3-Hydroxyacyl-CoA

PhaG

Thioesterase (TesA)

3-Ketoacyl-CoA

FabI Fatty acid 3-Ketoacyl-ACP biosynthesis pathway FabH FabA FabG

3-Hydroxyacyl-ACP

Enoyl-ACP

Acyl-ACP

Figure 4.2  Metabolic engineering strategies for the various PHAs from inherent intermediates of E. coli and Salmonella. Abbreviations: Buk, butyrate kinase from Clostridium acetobutylicum; FadB, enoyl-CoA hydratase; FadE, acyl-CoA dehydrogenase; GabT, glutamate:succinic semialdehyde transaminase from E. coli; GadA, glutamate decarboxylase from Escherichia coli; 4HbD, hydroxybutyrate dehydrogenase from Ralstonia eutropha; SucD, succinic semialdehyde dehydrogenase; PhaJ, (R)-specific enoyl-CoA hydratase; PrpE, propionyl-CoA synthetase from Salmonella enterica serovar typhimurium; Pta, phosphotransacetylase; Ptb, phosphotransbutyrylase from C. acetobutylicum.

3HV-CoA

Propionyl-CoA Acetyl-CoA

(2R)-methyl-malonyl-CoA

Succinyl-CoA

TCA-cycle α–ketoglutarate

Oxaloacetate

Acetyl-CoA

Pyruvate

Glyceraldehyde 3-phosphate

Glucose-P

4-10 Cellular Metabolism

Polymerization of Building Blocks to Macromolecules

4-11

4HB-CoA could be produced from succinyl-CoA in recombinant E. coli by three enzymes, including succinic semialdehyde dehydrogenase (SucD), 4-hydroxybutyrate dehydrogenase (4hbD), and 4-hydroxybutyryl-CoA: CoA transferase (Cat2) from Clostridium kluyveri [44]. By employing this pathway, recombinant E. coli could supply 4HB monomers from glucose to P(3HB-co-4HB) by up to 2.8 mol%. To construct a metabolic pathway in E. coli to generate 4HB from glutamate, glutamate decarboxylases from Arabidopsis thaliana or E. coli, glutamate:succinic semialdehyde transaminase from E. coli, 4-hydroxybutyrate dehydrogenase from R. eutropha, and 4-hydroxybutyrate-CoA: CoA transferase from C. kluyveri have been employed [45]. When the R. eutropha PHA biosynthesis genes were coexpressed with these engineered pathways, the copolymer of 3HB and 4HB was produced. However, the extent of 4HB assimilation into the copolymer was low (up to 3.5 mol%). 3HV-CoA, the substrate for the synthesis of PHAs containing 3HV, for example, P(3HB-co-3HV), is synthesized through condensation of propionyl-CoA with acetyl-CoA by β-ketothiolase (PhaA). Usually, propionyl-CoA is generated by activation of propionic acid in the medium. Mole fraction of 3HV in the copolymer is dependent on the amount of propionic acid added to the cells, but the concentration of propionic acid should be maintained in optimized level to support good cell growth along with desired 3HV composition due to the toxic effect of propionic acid at excess concentration. Development of metabolic pathway to generate propionyl-CoA from inherent intermediates of host strain can be a good solution to relieve toxicity of propionic acid during the synthesis of PHAs containing 3HV [46]. This was demonstrated by the expression of E. coli sbm and ygfG genes, which encode a novel (2R)-methylmalonylCoA mutase and a (2R)-methylmalonyl-CoA decarboxylase, respectively in prpC strain of Salmonella enterica serovar Typhimurium. Recombinant S. enterica expressing sbm and ygfG genes could convert succinyl-CoA, derived from the tricarboxylic acid cycle, to propionyl-CoA, and synthesized P(3HB-co3HV) from glycerol. It was possible to increase the 3HV fraction in the copolymer up to 15 mol%, which is usually obtainable by direct addition of propionic acid to the medium. To date, there is no report about the synthesis pathway of PHA containing 3HP, in which 3HP-CoA is generated from the inherent intermediates of host strain. Recently, Cargill proposed 3HP synthesis pathway via 3HP-CoA from glucose opening the possibility to produce PHA containing 3HP using metabolic intermediates of host strain [47]. Three metabolic pathways for the production of 3HP from glucose are proposed. Firstly, enzymes used during fermentation of alanine in Clostridium propionicum are employed. Lactate is activated to lactyl-CoA by CoA synthetase or CoA transferase and then converted to acrylyl-CoA by lactyl-CoA dehydratase. 3-hydroxypropionyl-CoA dehydratase adds one molecule of water into acrylyl-CoA to make 3HP-CoA. And then, 3HP-CoA is hydrolyzed by CoA hydrolase to release 3HP. Secondly, enzymes involved in the 3-hydroxypropionate cycle of autotrophic CO2 fixation in the phototrophic green non-sulfur bacterium Chloroflexus aurantiacus are used. In this pathway, malonyl-CoA is directly converted into 3HP by bifunctional enzyme malonyl-CoA reductase. Finally, a pathway, in which β-alanine is converted into 3HP via malonate semialdehyde by 4,4-aminobutyrate aminotransferase and 3-HP dehydrogenase is proposed. At present, only small amounts of 3HP can be produced using these pathways.

4.4.2  MCL-PHA and SCL-MCL-PHA MCL-PHA monomers are mainly provided from the intermediates of fatty acid metabolism because such metabolites can be easily converted into (R)-3-hydroxyacyl-CoAs (R3HA-CoAs), the most favorable substrates for PHA synthase. Enoyl-CoA, 3-ketoacyl-CoA, (S)-3-hydroxyacyl-CoA, and 3-hydroxyacyl-ACP are the major precursors of PHAs (Figure 4.2). In natural PHA producing bacteria, MCL-PHAs are efficiently synthesized from the intermediates of fatty acid β-oxidation and de novo fatty acid biosynthesis pathways. However, in recombinant E. coli, efficient production of PHAs with precise control of monomer composition using fatty acid metabolism has not yet been achieved because metabolic pathways involved in fatty acid metabolism are not optimized for the PHA synthesis. Enzymes supplying R3HA-CoAs from these fatty acid metabolism intermediates have

4-12

Cellular Metabolism

been identified and cloned from natural PHA-producing bacteria. These enzymes include (R)-specific enoyl-CoA hydratase (PhaJ) [48–53], 3-ketoacyl-ACP reductase (FabG) [54–56] and 3-hydroxydecanoyl-ACP:CoA transferase (PhaG) [57–60]. These enzymes have been characterized in molecular level and employed to develop PHA biosynthesis pathways in recombinant E. coli using intermediates of fatty acid metabolism. There is no specific metabolic engineering strategy for leading natural PHA producing bacteria to produce PHA from intermediates of fatty acid metabolism. When nutrient limitation is employed to the cell, bacteria naturally optimize their physiological state to accumulate PHA. Because E. coli is not able to synthesize PHA naturally, it is necessary to amplify specific genes directly involved in PHA biosynthesis obtained from natural PHA producing bacteria and to further create metabolic condition favorable to PHA biosynthesis through genetic and environmental modification. Engineering β-oxidation pathway and de novo fatty acid biosynthesis pathways strictly follows this strategy to support PHA biosynthesis. First, amplification of enzymes that directly convert intermediates of fatty acid metabolism to R3HACoAs, such as enoyl-CoA hydratase (PhaJ), 3-ketoacyl-ACP reductase (FabG), and 3-hydroxydecanoylACP: CoA transferase (PhaG) has been employed to construct PHA biosynthesis pathway. Presence of (R)-specific enoyl-CoA hydratase was revealed during characterization of PHA biosynthesis genes of A. caviae that is able to accumulate PHA consisting of 3HB and 3HHx from oils. In A. caviae, phaP, phaC, and phaJ, which encodes phasin, PHA synthase, enoyl-CoA hydratase, respectively, consist of the PHA biosynthesis operon. Based on the nucleotide sequence of A. caviae phaJ gene, various (R)-specific enoyl-CoA hydratases having different substrate specificities have been cloned and characterized from E. coli and other microorganisms which are able to produce PHA from fatty acid [48–53,61]. The basic idea that 3-ketoacyl-ACP reductase can mediate PHA biosynthesis using intermediates of β-oxidation pathway, 3-ketoacyl-CoA, was originated from the amino acid sequence homology of 3-ketoacyl-ACP reductase to acetoacetyl-CoA reductase (PhaB) [54,56]. Amplification of 3-ketoacyl-ACP reductases from E. coli and P. aeruginosa allowed us to develop MCL-PHA biosynthesis pathway from fatty acids. Because enoyl-CoA hydratase and 3-ketoacyl-ACP reductase compete with inherent β-oxidation pathway enzymes for intermediates, it was suggested that high-level expression of these genes is necessary to construct the metabolic link between β-oxidation and PHA biosynthetic pathways [62]. Pseudomonads belonging to rRNA homology group I synthesize MCL-PHA from glucose, indicating the de novo fatty acid biosynthesis pathway is connected to the PHA biosynthesis pathway. Recently, the phaG gene encoding 3-hydroxydecanoyl-ACP: CoA transferase was cloned from several Pseudomonas sp. and characterized at a molecular level for their role of constructing metabolic link between PHA biosynthesis pathway and fatty acid biosynthesis pathway [57–60]. However, expression of the phaG gene alone could not support MCL-PHA production from glucose in recombinant E. coli [57,63]. Therefore, E. coli FabH that might have 3-ketoacyl-ACP:CoA transferase activity, was examined for its activity to develop PHA biosynthesis pathway using intermediates of fatty acid biosynthesis pathway [64]. It was found that E. coli FabH has narrow activity to synthesize only 3HB-CoA. To broaden the substrate specificity of E. coli FabH, protein engineering of E. coli FabH was carried out by site-directed mutagenesis of Phe87 amino acid through in vitro saturated mutagenesis by PCR [65]. Phe87 amino acid was chosen as a target for mutagenesis on the basis of the crystal structure of Mycobacterium tuberculosis FabH that has specificity for substrates with C10–C16 carbon atoms. Several FabH mutant showed broader substrate specificity to generate R3HA-CoA having carbon atoms up to C12. The MCLPHA content obtained by expression of wild and engineered E. coli FabH was quite low (< 1%) and even Pseudomonas PhaG could not support PHA biosynthesis. This can be due to that the fatty acid biosynthesis pathway in E. coli was unable to supply enough PHA precursors, unlike the pathway in pseudomonads. Because degradation of fatty acid through β-oxidation is processed by multifunctional enzyme complex, FadBA, in which FadB has function of enoyl-CoA hydratase, 3-hydroxyacyl-CoA dehydrogenase, and 3-hydroxyacyl-CoA epimerase and FadA is 3-ketoacyl-CoA thiolase, major precursors for PHA biosynthesis, enoyl-CoA, (S)-3-hydroxyacyl-CoA, and 3-ketoacyl-CoA, are not easily accessible to PHA

Polymerization of Building Blocks to Macromolecules

4-13

biosynthesis pathway in E. coli that actively operates β-oxidation. Therefore, secondly, inhibition of enzymes involved in fatty acid metabolism has been tried to develop PHA biosynthesis pathway from β-oxidation. This strategy was further employed to construct PHA biosynthesis pathway using de novo fatty acid biosynthesis pathway. FadB and/or FadA mutant E. coli was found to be good hosts to synthesize MCL-PHA from fatty acid when MCL-PHA synthase was expressed [66,67]. Also, chemical inhibition of FadA by acrylic acid resulted in MCL-PHA biosynthesis [68]. When we examine the monomer composition of MCL-PHA obtained from E. coli deficient in β-oxidation, monomers having the same carbon numbers or numbers reduced by 2 and 4 or increased by 2 compared with those of the supplied fatty acid find to be incorporated into MCL-PHA [66]. For example, MCL-PHA consisting of 2.5 mol% 3-hydroxyhexanoate (3HHx), 20 mol% 3-hydroxyoctanoate (3HO), 72.5 mol% 3-hydroxydecanoate (3HD) and 5 mol% 3-hydroxydodecanoate (3HDD) was synthesized in fadB mutant E. coli expressing P. aeruginosa phaC1 gene from sodium decanoate [66]. Based on these results, it can be assumed that enigmatic enzyme substitutes for FadB to actively generate R3HA-CoA and to degrade fatty acid by two carbon atoms in fadB mutant E. coli strain. Molecular characterization of what enzymes are involved in the generation of R3HA-CoA in the absence of FadB and/or FadA found that various enzymes including YfcX [69], enoyl-CoA hydratase MaoC [61], and crotonase superfamily enzyme such as YdbU, PaaG, and PaaF [70] operate to convert enoyl-CoA to R3HA-CoA in fadB mutant E. coli. Deletion of genes encoding YfcX and MaoC in the chromosome of fadB mutant E. coli significantly decreased PHA synthesis activity from fatty acid. fadB yfcX and fadB maoC double mutant E. coli successfully recovered PHA biosynthesis activity from fatty acid by restoration of YfcX and MaoC in plasmid-based expression [61,69]. YfcX and YfcY homologous to FadB and FadA, respectively, were found to be components of another β-oxidation pathway operating under anaerobic conditions using nitrate as a terminal respiratory electron acceptor [71]. These enzymes proved to function normally in mutant strains lacking aerobic β-oxidation pathway enzymes. The substrate specificities of YfcXY are similar to those of FadBA. Based on these results, it can be concluded that YfcX is involved in PHA biosynthesis and degradation of acyl-CoAs in fadB mutant E. coli. Differences of YfcX, crotonase superfamily, and enoyl-CoA hydratase MaoC can be found in the PHA biosynthesis activities obtained by amplification of these enzymes in normal and fadB mutant E. coli strains. When YfcX and crotonase superfamily enzyme were amplified in normal E. coli strain, no PHA biosynthesis was achieved from fatty acid [61,70]. MCL-PHA content was increased without much modification of monomer composition of PHA when these enzymes were amplified in fadB mutant E. coli [61,70]. However, MaoC amplification in normal E. coli strain resulted in the accumulation of MCL-PHA from fatty acid and in fadB mutant E. coli strain, PHA with altered monomer composition was synthesized from fatty acid [61]. Monomer composition seems to be altered following substrate specificity of MaoC. Similar results that can be used for the comparison of enzymes involved in PHA biosynthesis in fadB mutant E. coli were obtained using P. oleovorans FadBA protein [53] and other (R)-specific enoyl-CoA hydratases [62]. As discussed above, even though it is necessary to achieve high-level expression of (R)-specific enoyl-CoA hydratases, PHA can be synthesized from fatty acid by additional expression of (R)-specific enoyl-CoA hydratases in normal E. coli strain that has functional β-oxidation pathway. However, P. oleovorans FadBA was not able to construct PHA biosynthesis pathway in normal E. coli strain even though FadBA was amplified by plasmid-based expression system. Considering the characteristic of FadBA, multifunctional enzyme complex, this seems to be due to not that FadBA do not dominantly perform hydration of enoyl-CoA in (R)-specific manner, but that they do not release R3HA-CoA making PHA synthase hardly accessible to R3HA-CoA during β-oxidation process of fatty acid. Strategy to mediate PHA biosynthesis by inhibition of enzymes related in fatty acid β-oxidation was also employed in construction of metabolic link between PHA biosynthesis and fatty acid biosynthesis. Chemicals that specifically inhibit the enzymes involved in fatty acid biosynthesis were examined for their ability to supply PHA precursors such as cerulenin for FabB and FabF and triclosan for FabI [63]. Only triclosan led to MCL-PHA synthesis from gluconate up to 2–3% dry cell in recombinant E. coli

4-14

Cellular Metabolism

expressing the P. putida phaG and the P. aeruginosa phaC1 genes. Unlike engineering of β-oxidation, fatty acid biosynthesis of E. coli could not successfully support MCL-PHA biosynthesis. It was suggested that weak metabolic activity of E. coli to generate intermediates of fatty acid biosynthesis is a factor for weak PHA biosynthesis activity based on the report that expression of the phaG gene only did not mediate MCL-PHA biosynthesis from glucose and E. coli could synthesize very small amounts of MCLPHA by expressing the phaG gene along with chemical inhibition via fatty acid biosynthesis pathway. However, recently, it was reported that E. coli expressing the phaG gene without PHA synthase efficiently excreted 3-hydroxydecanoic acid generated from the intermediates of fatty acid biosynthesis to the growth medium [72–74]. Addition of triclosan increased 3-hydroxydecanoic acid production by 20–30%. When the amount of excreted 3-hydroxydecanoic acid was considered on the basis of cell concentration, it reached nearly 50% of cell concentration [73]. This is quite interesting result to show that E. coli has sufficient metabolic capacity to supply enough fatty acid biosynthesis pathway intermediates for PHA biosynthesis. Thioesterase II (TesB) of E. coli might be reason to inhibit MCLPHA biosynthesis using intermediates of fatty acid biosynthesis through fast removal of CoA from R3HA-CoA, resulting in the excretion of overproduced (R)-3-hydroxyalkanoic acid [74]. When the tesB mutant E. coli strain was employed for the excretory production of 3-hydroxydecanoic acid from glucose by expressing the phaG gene, production of 3-hydroxydecanoic acid was decreased to 72 mg/L that is only 10% of 3-hydroxydecanoic acid concentration achieved with control strain. It was revealed that expression of the tesB gene was triggered by accumulation of 3HD-CoA that is synthesized by PhaG. The exact role of TesB in MCL-PHA biosynthesis using intermediates of fatty acid biosynthesis will be resolved by the experiment about the production of MCL-PHA in the tesB mutant E. coli by expression of the phaG gene and the phaC gene. Interestingly, thioesterase has been employed to link fatty acid biosynthesis and fatty acid β-oxidation pathway to construct MCL-PHA biosynthesis pathway from glucose in the fad mutant E. coli that constitutively operates impaired β-oxidation pathway [75,76]. fadR fadB mutant E. coli could produce MCL-PHA from gluconate up to 2.3% dry cell weight by expressing E. coli thioesterase I and P. oleovorans PHA synthase [75]. Also, expression of acyl-ACP thioesterase from the plant Umbellularia californica established MCL-PHA biosynthesis from gluconate in recombinant E. coli [76]. Accumulated laurate by U. californica acyl-ACP thioesterase was converted to PHA precursors through the β-oxidation pathway in fadB mutant E. coli, and then polymerized into MCL-PHA by P. aeruginosa MCL-PHA synthase. Based on these results of MCL-PHA biosynthesis obtained by employing thioesterase, we can suggest two possible strategies to produce MCL-PHA from glucose in recombinant E. coli: (I) amplification of the phaG and phaC genes in the tesB mutant E. coli strain, (II) amplification of the phaG, tesB, and phaC genes in fadR fadA and/or fadB mutant.

4.5  Conclusion PHA biosynthesis is a good model system for examining the metabolic engineering strategies for efficient biopolymer production. As reviewed in this chapter, successful metabolic engineering of microorganisms for the enhanced production of biopolymers and/or production of novel biopolymers requires a priori knowledge. Detailed understanding on the metabolic pathways leading to the synthesis of a biopolymer as well as the genes and enzymes involved is essential. Even though successful metabolic engineering can be performed using this information, it will be better to optimize the upstream to downstream processes at the same time; metabolic engineering of strains should be performed after considering fermentation and recovery/purification processes. As biopolymers like PHAs should be relatively inexpensive, the entire process should be optimized with economical production in mind. Carbon substrate can be a cost-determining factor in the production of bulk biopolymers, and thus the production strain should be selected or engineered to utilize an inexpensive carbon source available at the production site. Recent advances in omics and systems biology disciplines are opening a new possibility of systemic optimization of strains and bioprocesses. Systems biotechnology [77,78] by

Polymerization of Building Blocks to Macromolecules

4-15

integrating genome, transcriptome, proteome, metabolome, and computational analyses are allowing us to identify new targets and new strategies for the metabolic engineering of microorganisms for the overproduction of bioproducts and production of novel products. This is also true for the production of biopolymers. We are observing the rapid rise of oil prices, which is reminding us of the finite nature of fossil fuels. Various chemicals and polymers we are using everyday need to be produced from renewable resources in the future. As the efficiencies of bioprocesses for the production of these chemicals and biopolymers by microbial fermentation are generally low, we need to improve the performance of the strains to be employed. Metabolic engineering strategies described in this chapter for PHA as an example should be useful for designing strains for other biopolymers. Furthermore, it is expected that the better metabolic engineering strategies will become available through the systems biotechnology.

Acknowledgments Our work described in this chapter was supported by the Korean Systems Biology Research Program (M10309020000-03B5002-00000) of the Ministry of Science and Technology. Further supports by the LG Chem Chair Professorship and KOSEF through the CUPS are appreciated.

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15. Fukui, T. et al. Biosynthesis of poly(3-hydroxybutyrate-co-3-hydroxyvalerate-co-3-hydroxy-heptanoate) terpolymers by recombinant Alcaligenes eutrophus. Biotechnol. Lett., 19, 1093, 1997. 16. Dennis, D. et al. Formation of poly(3-hydroxybutyrate-co-3-hydroxyhexanoate) by PHA synthase from Ralstonia eutropha. J. Biotechnol., 64, 177, 1998. 17. Antonio, R.V., Steinbüchel, A., and Rehm, B.H.A. Analysis of in vivo substrate specificity of the PHA synthase from Ralstonia eutropha: formation of novel copolyesters in recombinant Escherichia coli. FEMS Microbiol. Lett., 182, 111, 2000. 18. Jia, Y. et al. Lipases provide a new mechanistic model for polyhydroxybutyrate (PHB) synthases: characterization of the functional residues in Chromatium vinosum PHB synthase. Biochemistry, 39, 3927, 2000. 19. Takase, K. et al. Alteration of substrate chain-length specificity of type II synthase for polyhydroxyalkanoate biosynthesis by in vitro evolution: in vivo and in vitro enzyme assays. Biomacromolecules, 5, 480, 2004. 20. Normi, Y.M. et al. Site-directed saturation mutagenesis at residue F420 and recombination with another beneficial mutation of Ralstonia eutropha polyhydroxyalkanoate synthase. Biotechnol. Lett. 27, 705, 2005. 21. Normi, Y.M. et al. Characterization and properties of G4X mutants of Ralstonia eutropha PHA synthase for poly(3-hydroxybutyrate) biosynthesis in Escherichia coli. Macromol. Biosci., 5, 197, 2005. 22. Matsumoto, K. et al. Synergistic effects of Glu130Asp substitution in the type II polyhydroxyalkanoate (PHA) synthase: enhancement of PHA production and alteration of polymer molecular weight. Biomacromolecules, 6, 99, 2005. 23. Takase, K. et al. Alteration of substrate chain-length specificity of type II synthase for polyhydroxyalkanoate biosynthesis by in vitro evolution: in vivo and in vitro enzyme assays. Biomacromolecules, 5, 480, 2004. 24. Takase, K., Taguchi, S., and Doi, Y. Enhanced synthesis of poly(3-hydroxybutyrate) in recombinant Escherichia coli by means of error-prone PCR mutagenesis, saturation mutagenesis, and in vitro recombination of the type II polyhydroxyalkanoate synthase gene. J. Biochem. (Tokyo), 133, 139, 2003. 25. Taguchi, S. et al. In vitro evolution of a polyhydroxybutyrate synthase by intragenic suppressiontype mutagenesis. J. Biochem. (Tokyo), 131, 801, 2002. 26. Kichise, T., Taguchi, S., and Doi, Y. Enhanced accumulation and changed monomer composition in polyhydroxyalkanoate (PHA) copolyester by in vitro evolution of Aeromonas caviae PHA synthase. Appl. Environ. Microbiol., 68, 2411, 2002. 27. Taguchi, S. et al. Analysis of mutational effects of a polyhydroxybutyrate (PHB) polymerase on bacterial PHB accumulation using an in vivo assay system. FEMS Microbiol. Lett., 198, 65, 2001. 28. Rehm, B.H.A. et al. Molecular characterization of the poly(3-hydroxybutyrate) (PHB) synthase from Ralstonia eutropha: in vitro evolution, site-specific mutagenesis and development of a PHB synthase protein model. Biochim. Biophys. Acta., 1594, 178, 2002. 29. Amara, A.A., Steinbüchel, A., and Rehm, B.H.A. In vivo evolution of the Aeromonas punctata polyhydroxyalkanoate (PHA) synthase: isolation and characterization of modified PHA synthases with enhanced activity. Appl. Microbiol. Biotechnol., 59, 477, 2002. 30. Amara, A.A. and Rehm, B.H.A. Replacement of the catalytic nucleophile cysteine-296 by serine in class II polyhydroxyalkanoate synthase from Pseudomonas aeruginosa mediated synthesis of a new polyester: identification of catalytic residues. Biochem. J., 374, 413, 2003. 31. Sheu, D.S. and Lee, C.Y. Altering the substrate specificity of polyhydroxyalkanoate synthase 1 derived from Pseudomonas putida GPo1 by localized semirandom mutagenesis. J. Bacteriol., 186, 4177, 2004. 32. Hein, S. et al. Biosynthesis of poly(4-hydroxybutyric acid) by recombinant strains of Escherichia coli. FEMS Microbiol. Lett., 153, 411, 1997.

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33. Liu, S.J. and Steinbüchel, A. A novel genetically engineered pathway for synthesis of poly(hydroxyalkanoic acids) in Escherichia coli. Appl. Environ. Microbiol., 66, 739, 2000. 34. Valentin, H.E. et al. Application of a propionyl coenzyme A synthetase for poly(3-hydroxypropionate-co-3-hydroxybutyrate) accumulation in recombinant Escherichia coli. Appl. Environ. Microbiol., 66, 5253, 2000. 35. Lim, S.J. et al. Amplification of the NADPH-related genes zwf and gnd for the oddball biosynthesis of PHB in an E. coli transformant harboring a cloned phbCAB operon. J. Biosci. Bioeng., 93, 543, 2002. 36. Lee, S.Y. Bacterial polyhydroxyalkanoates. Biotechnol. Bioeng., 49, 1, 1996. 37. Lee, S.Y. Plastic bacteria? Progress and prospects for polyhydroxyalkanoate production in bacteria. Trends Biotechnol., 14, 431, 1996 38. Madison, L.L. and Huisman, G.W. Metabolic engineering of poly(3-hydroxyalkanoates): from DNA to plastic. Microbiol. Mol. Biol. Rev., 63, 21, 1999. 39. Han, M.J., Yoon, S.S., and Lee, S.Y. Proteome analysis of metabolically engineered Escherichia coli producing poly(3-hydroxybutyrate). J. Bacteriol., 183, 301, 2001. 40. Hong, S.H. et al. In silico prediction and validation of the importance of Entner-Doudoroff pathway in poly(3-hydroxybutyrate) production by metabolically engineered Escherichia coli. Biotech. Bioeng., 83, 854, 2003. 41. Han, M.-J. et al. Analysis of poly(3-hydroxybutyrate) granule-associated proteome in recombinant Escherichia coli. J. Microbiol. Biotechnol., 16, 901, 2006. 42. Wong, H.H. et al. Metabolic analysis of poly(3-hydroxybutyrate) production by recombinant Escherichia coli. J. Microbiol. Biotechnol., 9, 593, 1999. 43. Van Wegen, R.J., Lee, S.Y., and Middelberg, A. Metabolic and kinetic analysis of poly(3-hydroxybutyrate) production by recombinant Escherichia coli. Biotechnol. Bioeng., 74, 70, 2001. 44. Valentin, H.E. and Dennis, D. Production of poly(3-hydroxybutyrate-co-4-hydroxybutyrate) in recombinant Escherichia coli grown on glucose. J. Biotechnol., 58, 33, 1997. 45. Valentin, H.E., Reiser, S., and Gruys, K.J. Poly(3-hydroxybutyrate-co-4-hydroxybutyrate) formation from gamma-aminobutyrate and glutamate. Biotechnol. Bioeng., 67, 291, 2000. 46. Aldor, I.S. et al. Metabolic engineering of a novel propionate-independent pathway for the production of poly(3-hydroxybutyrate-co-3-hydroxyvalerate) in recombinant Salmonella enterica serovar typhimurium. Appl. Environ. Microbiol., 68, 3848, 2002. 47. Gokarn, R.R. et al. PCT patent WO0242418. 48. Fukui, T. and Doi, Y. Expression and characterization of (R)-specific enoyl Coenzyme A hydratase involved in polyhydroxyalkanoate biosynthesis by Aeromonas caviae. J. Bacteriol., 180, 667, 1998. 49. Tsuge, T. et al. Molecular cloning of two (R)-specific enoyl-CoA hydratase genes from Pseudomonas aeruginosa and their use for polyhydroxyalkanoate synthesis. FEMS Microbiol. Lett., 184, 193, 2000. 50. Tsuge, T. et al. Molecular characterization and properties of (R)-specific enoyl-CoA hydratases from Pseudomonas aeruginosa: metabolic tools for synthesis of polyhydroxyalkanoates via fatty acid β-oxidation. Int. J. Biol. Macromol., 31, 195, 2003. 51. Park, S.J. et al. Production of poly(3-hydroxybutyrate-co-3-hydroxyhexanoate) by metabolically engineered Escherichia coli strains. Biomacromolecules, 2, 248, 2001. 52. Tsuge, T. et al. Alteration of chain length substrate specificity of Aeromonas caviae R-enantiomerspecific enoyl-coenzyme A hydratase through site-directed mutagenesis. Appl. Environ. Microbiol., 69, 4830, 2003. 53. Fiedler, S., Steinbüchel, A., and Rehm, B.H.A. The role of the fatty acid beta-oxidation multienzyme complex from Pseudomonas oleovorans in polyhydroxyalkanoate biosynthesis: molecular characterization of the fadBA operon from P. oleovorans and of the enoyl-CoA hydratase genes phaJ from P. oleovorans and Pseudomonas putida. Arch. Microbiol., 178, 149, 2002.

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54. Ren, Q. et al. FabG, an NADPH-dependent 3-ketoacyl reductase of Pseudomonas aeruginosa, provides precursors for medium-chain-length poly-3-hydroxyalkanoate biosynthesis in Escherichia coli. J. Bacteriol., 182, 2978, 2000. 55. Park, S.J., Park, J.P., and Lee, S.Y. Metabolic engineering of Escherichia coli for the production of medium-chain-length polyhydroxyalkanoates rich in specific monomers. FEMS Microbiol. Lett., 214, 217, 2002. 56. Taguchi, K. et al. Co-expression of 3-ketoacyl-ACP reductase and polyhydroxyalkanoate synthase genes induces PHA production in Escherichia coli HB101 strain. FEMS Microbiol. Lett., 176, 183, 1999. 57. Rehm, B.H.A., Kruger, N., and Steinbüchel, A. A new metabolic link between fatty acid de novo synthesis and polyhydroxyalkanoic acid synthesis. The phaG gene from Pseudomonas putida KT2440 encodes a 3-hydroxyacyl-acyl carrier protein-coenzyme a transferase. J. Biol. Chem., 273, 24044, 1998. 58. Hoffmann, N., Steinbüchel, A., and Rehm, B.H.A. The Pseudomonas aeruginosa phaG gene product is involved in the synthesis of polyhydroxyalkanoic acid consisting of medium-chain-length constituents from non-related carbon sources. FEMS Microbiol. Lett., 184, 253, 2000. 59. Hoffmann, N., Steinbüchel, A., and Rehm, B.H.A. Homologous functional expression of cryptic phaG from Pseudomonas olevorans establishes the transacylase-mediated polyhydroxyalkanoate biosynthetic pathway. Appl. Microbiol. Biotechnol., 54, 665, 2000. 60. Matsumoto, K. et al. Cloning and characterization of the Pseudomonas sp. 61-3 phaG gene involved in polyhydroxyalkanoate biosynthesis. Biomacromolecules, 2, 142, 2001. 61. Park, S.J. and Lee, S.Y. Identification and characterization of a new enoyl coenzyme A hydratase involved in biosynthesis of medium-chain-length polyhydroxyalkanoates in recombinant Escherichia coli. J. Bacteriol., 185, 5391, 2003. 62. Fukui, T. et al. Co-expression of polyhydroxyalkanoate synthase and (R)-enoyl-CoA hydratase genes of Aeromonas caviae establishes copolyester biosynthesis pathway in Escherichia coli. FEMS Microbiol. Lett., 170, 69, 1999. 63. Rehm, B.H.A., Mitsky, T.A., and Steinbüchel, A. Role of fatty acid de novo biosynthesis in polyhydroxyalkanoic acid (PHA) and rhamnolipid synthesis by pseudomonads: establishment of the transacylase (PhaG)-mediated pathway for PHA biosynthesis in Escherichia coli. Appl. Environ. Microbiol., 67, 3102, 2001. 64. Taguchi, K. et al. Over-expression of 3-ketoacyl-ACP synthase III or malonyl-CoA-ACP transacylase gene induces monomer supply for the polyhydroxybutyrate production in Escherichia coli HB101. Biotechnol. Lett., 21, 579, 1999. 65. Nomura, C.T. et al. Coexpression of genetically engineered 3-ketoacyl-ACP synthase III (fabH) and polyhydroxyalkanoate synthase (phaC) genes leads to short-chain-length-medium-chain-length polyhydroxyalkanoate copolymer production from glucose in Escherichia coli JM109. Appl. Environ. Microbiol., 70, 999, 2004. 66. Langenbach, S., Rehm, B.H.A., and Steinbüchel, A. Functional expression of the PHA synthase gene phaC1 from Pseudomonas aeruginosa in Escherichia coli results in poly(3-hydroxyalkanoate) synthesis. FEMS Microbiol. Lett., 150, 303, 1997. 67. Qi, Q., Rehm, B.H.A., and Steinbüchel, A. Synthesis of poly(3-hydroxyalkanoates) in Escherichia coli expressing the PHA synthase gene phaC2 from Pseudomonas aeruginosa: comparison of PhaC1 and PhaC2. FEMS Microbiol. Lett., 157, 155, 1997. 68. Qi, Q., Rehm, B.H.A., and Steinbüchel, A. Metabolic routing towards polyhydroxyalkanoic acid synthesis in recombinant Escherichia coli (fadR): inhibition of fatty acid beta-oxidation by acrylic acid. FEMS Microbiol. Lett., 167, 89, 1998. 69. Snell, K.D. et al. YfcX enables medium-chain-length poly(3-hydroxyalkanoate) formation from fatty acids in recombinant Escherichia coli fadB strains. J. Bacteriol., 184, 5696, 2002.

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70. Park, S.J. and Lee, S.Y. New FadB homologous enzymes and their use in enhanced biosynthesis of medium-chain-length polyhydroxyalkanoates in fadB mutant Escherichia coli. Biotechnol. Bioeng., 86, 681, 2004. 71. Campbell, J.W., Morgan-Kiss, R.M., and Cronan, J.E., Jr. A new Escherichia coli metabolic competency: growth on fatty acids by a novel anaerobic β-oxidation pathway. Mol. Microbiol., 47, 793, 2003. 72. Zhao, K., et al. Production of D-(-)-3-hydroxyalkanoic acid by recombinant Escherichia coli. FEMS Microbiol. Lett., 218, 59, 2003. 73. Zheng, Z. et al. Production of 3-hydroxydecanoic acid by recombinant Escherichia coli HB101 harboring phaG gene. Antonie Van Leeuwenhoek., 85, 93, 2004. 74. Zheng, Z. et al. Thioesterase II of Escherichia coli plays an important role in 3-hydroxydecanoic acid production. Appl. Environ. Microbiol., 70, 3807, 2004. 75. Klinke, S. et al. Production of medium-chain-length poly(3-hydroxyalkanoates) from gluconate by recombinant Escherichia coli. Appl. Environ. Microbiol., 65, 540, 1999. 76. Rehm, B.H.A. and Steinbüchel, A. Heterologous expression of the acyl-acyl carrier protein thioesterase gene from the plant Umbellularia californica mediates polyhydroxyalkanoate biosynthesis in recombinant Escherichia coli. Appl. Microbiol. Biotechnol., 55, 205, 2001. 77. Lee, S.Y., Lee, D.Y., and Kim, T.Y. Systems biotechnology for strain improvement. Trends Biotechnol. 23, 331, 2005. 78. Park, S.J. et al. Global physiological understanding and metabolic engineering of microorganisms based on omics studies. Appl. Microbiol. Biotechnol. 68, 567, 2005.

5 Rare Metabolic Conversions—Harvesting Diversity through Nature 5.1 5.2 5.3 5.4

Manuel Ferrer Institute of Catalysis

Peter N. Golyshin HZI-Helmholtz Centre for Infection Research

Introduction ���������������������������������������������������������������������������������������5-1 How Diverse Are Functional Groups?........................................... 5-4 Diversity of Enyzmes and Current Frontiers for Bioconversions ���������������������������������������������������������������������������������� 5-5 Main Chemical Conversions Mediated by Enzymes: Putative “Rare” Conversions ���������������������������������������������������������� 5-8

Hydrolytic Enzymes in Enantio-Transformations  •  Rare Conversions Using Carbohydrate Modifying Enzymes: Production of Rare Carbohydrates  •  Oxidative Enzymes for the Conversion of Organic Pollutants in Industrial Effluents to Useful Products  •  Rare Conversions Using Fatty Acids: Production of Unique Complex Lipids and Essential Fatty Acids  •  Rare Conversions Using Aliphatic Groups: Incorporation of Chlorine, Fluorine, Bromide, and Iodine to Peptides-Polyketides  •  Rare Conversions Using Amino Acids: Production of Unnatural Amino Acids  •  Rare Conversion with Flavonoids and Steroids: Drug Synthesis  •  Rare Conversion of Xenobiotics: Potential Biodegradation Pathways  •  Rare Routes for Synthesis of Hormones and Vitamins: Rare Carboxylations and Hydroxylations  •  Cofactor-Dependent Enzymes and Cofactor Regeneration  •  Enantiomerically Pure Compounds by Rare Domino and MCRs

5.5 5.6

How Can New Catalytic Functions Be Achieved?......................5-16  ecent Advances in Metagenomics: The Untapped R Reservoir of Proteins from Unculturable Microbes...................5-18 Acknowledgments ������������������������������������������������������������������������������������� 5-20 References ����������������������������������������������������������������������������������������������������5-21

5.1  Introduction At the fundamental level the cellular metabolism is mediated through the enzymes that catalyze biotransformation reactions. The specific reactions transforming the particular substrate in an experimental assay (or in an in vivo system) may be or may not be of a practical interest. However, in case the research focuses on biotransformations-conversions and their biotechnological applications, in most cases a knowledge of a whole genetic circuit, particular pathway, and individual enzymes comprising the pathway are needed. Traditionally, the screening for a specific biocatalytic reaction was performed through the growth of a particular microorganism and the disappearance of the starting compound (reaction substrate) and detection of its disappearance, or/and occurrence and accumulation of 5-1

5-2

Cellular Metabolism

corresponding metabolites [1,2]. The structures of metabolites and the stoichiometry of their formation (determined both, experimentally or in silico) give some important hints to the specificity of an enzyme and its probable usefulness for the application [3]. Whatever the case, the reactions of an apparent new pathway are routinely elucidated via molecular biological methods, typically through molecular cloning and expression of the protein in a surrogate host, such as Escherichia coli, and consequent characterization of its enzymatic activity (Table 5.1). To obtain in a desired biocatalyst a known pathway or reaction Table 5.1  Classification of Enzymes and EC Numbers 1.  Oxidoreductases    1.1  Acting on the CH-OH group of donors    1.2  Acting on the aldehyde or oxo group of donors    1.3  Acting on the CH-CH group of donors    1.4  Acting on the CH-NH2 group of donors    1.5  Acting on the CH-NH group of donors    1.6  Acting on NADH or NADPH    1.7  Acting on other nitrogenous compounds as donors    1.8  Acting on a sulfur group of donors    1.9  Acting on a heme group of donors 1.10  Acting on diphenols and related substances as donors 1.11  Acting on a peroxide as acceptor 1.12  Acting on hydrogen as donor 1.13  Acting on single donors with incorporation of molecular oxygen (oxygenases) 1.14  Acting on paired donors with incorporation of molecular oxygen 1.15  Acting on superoxide as acceptor 1.16  Oxidizing metal ions 1.17  Acting on CH or CH2 groups 1.18  Acting on iron-sulfur proteins as donors 1.19  Acting on reduced flavodoxin as donor 1.20  Acting on phosphorus or arsenic in donors 1.21  Acting on X-H and Y-H to form an X-Y bond 1.97  Other oxidoreductases 1.-  Other oxidoreductases 2.  Transferases 2.1  Transferring one-carbon groups 2.2  Transferring aldehyde or ketone residues 2.3  Acyltransferases 2.4  Glycosyltransferases 2.5  Transferring alkyl or aryl groups, other than methyl groups 2.6  Transferring nitrogenous groups 2.7  Transferring phosphorus-containing groups 2.8  Transferring sulfur-containing groups 2.9  Transferring selenium-containing groups 2.-  Transferring selenium-containing groups 3.  Hydrolases 3.1  Acting on ester bonds 3.2  Glycosidases 3.3  Acting on ether bonds 3.4  Acting on peptide bonds (peptidases) (Continued)

Rare Metabolic Conversions—Harvesting Diversity through Nature

5-3

Table 5.1  (Continued)    3.5  Acting on carbon–nitrogen bonds, other than peptide bonds    3.6  Acting on acid anhydrides    3.7  Acting on carbon–carbon bonds    3.8  Acting on halide bonds    3.9  Acting on phosphorus–nitrogen bonds 3.10  Acting on sulfur–nitrogen bonds 3.11  Acting on carbon–phosphorus bonds 3.12  Acting on sulfur–sulfur bonds 3.13  Acting on carbon–sulfur bonds 3.-  Acting on carbon–sulfur bonds 4.  Lyases 4.1  Carbon–carbon lyases 4.2  Carbon–oxygen lyases 4.3  Carbon–nitrogen lyases 4.4  Carbon–sulfur lyases 4.5  Carbon–halide lyases 4.6  Phosphorus–oxygen lyases 4.99  Other lyases 5.  Isomerases 5.1  Racemases and epimerases 5.2  cis-trans-Isomerases 5.3  Intramolecular oxidoreductases 5.4  Intramolecular transferases (mutases) 5.5  Intramolecular lyases 5.99  Other isomerases 5.-  Other isomerases 6.  Ligases 6.1  Forming carbon–oxygen bonds 6.2  Forming carbon–sulfur bonds 6.3  Forming carbon–nitrogen bonds 6.4  Forming carbon–carbon bonds 6.5  Forming phosphoric ester bonds 6.6  Forming nitrogen–metal bonds 6.-  Forming nitrogen–metal bonds

might be taken. The goal is then to obtain an ideal enzyme which is highly active with a given substrate or under specific set of conditions [4]. In this case, the screening method is important, as it might be necessary to look at hundreds or thousands of enzymes in the first rounds of screening. The description of a wide range of screening methods can be seen in numerous publications, the most relevant being those by the groups of Omura and Cutler [5,6]. Moreover, fast, efficient, and high-throughput screening methods that could test vast numbers of enzymes for specific applications have been discovered in the last decades, bringing made to order enzymes a step closer [7]. Recently, a screening system utilizing the “fluorescence activated cell sorting,” or FACS, a technology that enables the identification of biological activity within a single cell, has been developed by Diversa Co. (CA). This system incorporates a laser with multiple wavelength capabilities and the ability to screen up to 50,000 clones per second, or over 1 billion clones per day. Another approach is to test existing isolates of microorganisms to determine if they have an enzyme which show a high activity with the substrate of interest. The pure

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Cellular Metabolism

cultures of microorganisms have passed the test of time well: those have successfully been used for over 100 years. However, will the advent of the genome-wide high-throughput techniques of enzymatic screening and in silico predictions [8], make the cultivation-based enzyme discovery excessive? Based on genome sequencing data it is currently possible to make a metabolic reconstruction in which all possible known biochemical conversions could be identified. Thus, a comprehensive metabolic reconstruction on aspergilli revealed up to 14,400 ORFs, 335 reactions, and 284 metabolites distributed over the intra- and extracellular compartments [9]. Moreover, the genomes of extremophilic bacteria and archaea could be even more sophisticated: those from Thermotoga, Sulfolobus, Picrophilus spp., containing the ORF numbers in the range of 1,500–4,000, may comprise more complex biochemistries in terms of unique proteins, metabolites, and enzymes under extreme physical-chemical conditions existing in their natural environment [10–12]. Even though the valuable genomic information of individual organisms represents a good starting point, the cultivation of microbes often represents a bottleneck. The biological diversity on our planet is mostly comprised of microorganisms, in terms of both absolute numbers of cells and in terms of the biomass and the numbers of species [13]. Recently it has been reported that up to 25,000 different genotypes in a millilitre of the seawater sample in some marine ecosystems have been found [14]. However, the majority of microbes in nature will never be cultured, and to access this great inexhaustable genetic and metabolic diversity the new “metagenomic” approach is used [15–21]. This approach uses harvesting DNA from an environmental sample (or from an enrichment), its archiving in the metagenomic libraries in appropriate hosts, and screening these libraries for a gene of interest or expressing the DNA and screening for the enzymatic activities of interest [17]. Alternatively, these libraries are subjected to a high throughput shotgun sequencing and automated annotation, which often produces thousands of new genes whose functions often remain unknown. This has been shown recently by the group of Venter [22], whose shotgun sequencing of environmental DNA from Sargasso Sea yielded 1,045 billion base pairs derived from 1,800 genomic species and deduced more than 1.2 million of previously unknown genes. However, the work reflects difficulties on genome assembly and other challenges in interpretation of large data sets. Metagenomics may result in the development of a number of new products and to provide the tools for the resolution of rare conversions which are not amenable to the existing biocatalysts. Below we focus on the issues, what are current challenges for enzymatic conversions and how the natural biodiversity may help to deal with these challenges.

5.2  How Diverse Are Functional Groups? The compounds to be considered in the context of conversions are the known organic molecules, an ever-expanding set of over 10 million compounds, both natural and artificial, which are divided in a variety of chemical functional groups with a definable set of chemical reactivities. However, it is to be expected that in the natural biological world, a huge number of undiscovered natural products and therefore “exotic-rare” chemical reactions for their synthesis and/or conversion awaits discovery. This conclusion is supported by the enormous taxonomic diversity of microorganisms and thus the corresponding unmined riches of yet unexplored catalytic diversity [12]. Furthermore, for enzymes catalyzing sequential reactions in a biosynthetic pathway, new catalytic functions could evolve after gene duplication, whereas the binding capacity for a common ligand is retained and the chemistry of catalysis is changed [23,24]. Alternatively, for enzymes catalyzing chemically related reactions in different biosynthetic pathways, after gene duplication the chemistry of catalysis could be retained but the substrate specificity can change [25]. In addition, some divergently evolved enzymes could have been derived from a largely conserved active site architecture that was modified to catalyze different reactions which are unrelated both with respect to chemical mechanism and substrate structure [26,27]. As the number and diversity of enzymes is practically unlimited (in terms of amino acid combinations), there is important work to be done in categorizing the known and unknown reactions via a functional group approach, and experimental science, uncovering new enzymatic reactions useful for the biosynthesis and catabolism of previously unstudied functional groups.

5-5

Rare Metabolic Conversions—Harvesting Diversity through Nature

Recycling of materials and waste elimination

Products that improve our diet and health

Food/Feed Genetic variants of plants to resist plagues, and scarcity of water and nutrients and to improve yield Agriculture chemicals to improve yields

Biopolymers: replacement of tissues, organs, bones, etc.

Selective and promiscuous biocatalysts

Material science

Chemical synthesis

Preparation of chiral synthons New ways to produce known or new products: anti-cancer, etc.

Energy and sustainable development

Biotechnology

Terrorism issues

Hydrogenases and laccases more stable and less inhibited by CO

Detection of chemical weapons / biological agents (virus/toxins/infectious bacteria)

Bioenergy: biofuels, bioethanol, biofuel cells

Figure 5.1  Challenges for biotechnology and possible points for implementation of rare enzymes and rare conversions.

The bibliographic records of the biotechnology research of past few decades, especially regarding the new enzymatic conversions, show that this emerging field may provide a clear alternative of those processes to the chemical synthetic industrial methods. Biocatalysts show regio-, quimio-, and stero-selectivity as along with a high catalytic efficiency and offer a great diversity of applications. Although microbial cultivation generates huge collections of microorganisms and their genes, it has been demonstrated that laboratory protein engineering evolution technology with expression-based and high-throughput screening technologies used for the optimization of enzymes and other proteins, also generates large and potentially valuable gene libraries and enzyme variants useful for particular conversions [4]. However, some processes remain to be resolved (see examples in Figure 5.1), which will require new biocatalysts (Figure 5.2).

5.3 Diversity of Enyzmes and Current Frontiers for Bioconversions Like all catalysts, enzymes are catalytically versatile agents which accelerate the rates of reactions normally undergoing no permanent chemical modifications resulting in the course of catalysis. Enzymes can accelerate, often by several orders of magnitude, reactions that under the mild conditions of cellular concentrations, temperature, pH, and pressure would proceed imperceptibly (or not at all) in the absence of the enzyme. Consonant with their role as biological catalysts, enzymes show considerable selectivity for the molecules (substrates) upon which they act. This makes enzyme and whole-cell biocatalysts a greener alternative to traditional organic synthesis which offers appropriate tools for the industrial transformation of natural or synthetic materials under mild reaction conditions [28,29], low energy requirements and minimizes the problems of isomerization and rearrangement [30,31]. One of the key findings in the last 15 years was the discovery that the enzymes may be active in organic solvents, and that this influences enzyme properties [32,33]. This is of a special interest for enzymes such as penicillin acylases, esterases, lipases, proteases, nitrilases, hydantoinases, oxidoreductases, and

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Cellular Metabolism Reaction engineering: To modulate the microenvironment of the enzyme

Biocatalysts engineering– protein engineering

To use known biocatalysts and to try to improve their activities and properties

New enzymes? Better enzymes? To study the microbial diversity

Cultivation independent methods The microbial world is characterized by its extreme biochemical diversity with more than 10 million species and a considerable divergence of gene sequences among species A large diversity of microbes needs to be studied to gain sufficient understanding of their functional roles

Figure 5.2  Alternatives for engineering or finding new enzymes and conversions. Table 5.2  Enzymes Commonly Used in Chemical Conversions Enzymes Esterases, lipases Amidases, proteases, and acylases Dehydrogenases Oxidases (mono- and dioxygenase) Peroxidases Kinases Aldolases and transketolases Glycosidases and glycosyltransferases Phosphorylases and phosphatases Sulphotransferases Transaminases Hydrolases Isomerases, lyases, and hydratases

Conversion Ester hydrolysis and formation Amide hydrolysis and formation Oxireduction of alcohols and ketones Oxidation Oxidation, epoxidation, and halohydratation Phosphorylation Aldol reaction (C-C-bond) Glycosidic bond formation Formation and hydrolysis of phosphate Formation of sulphate esters Amino acid synthesis (C-N bond) Hydrolysis Isomerization, addition, elimination, and replacement

transferases, which have been successfully applied for the synthesis of numerous bioactive compounds [34]. However, most of the enzymes react with only a small group of closely related chemical compounds; many demonstrate absolute specificity, having only one substrate molecule which is appropriate for reaction (see Table 5.2 for specific conversions and classification of enzymes according to them). Over 3,000 enzymes have so far been identified, and this number may be greatly augmented in the wake of in silico data mining and proteomic and (meta)genomic research [20,34]. The last strategy is of a special interest as we know that the microbial word is characterized by its extreme biochemical diversity

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with more than 10 million species and a considerable divergence of gene sequences with and among species. Moreover, a large diversity of microbes need to be studied to gain sufficient understanding of their functional roles.Although the existing experimental and genome data have revealed a battery of enzymes whose physiological roles and biotechnological potential are well established in laboratory and industry, we strongly believe that the enzymology is still in its infancy and its future is going to greatly expand during the forthcoming search for novel and versatile enzymes able to catalyze reactions that are difficult to perform by chemical methods (Figure 5.2) [20]. Biotransformation processes are far more diverse than one can imagine simply counting all existing protein families [35]. Thousands of different strains and enzymes are required to exploit the selective biotransformation potential for the conversion of a myriad of different substrates into the desired products, especially new optically active derivatives. This may offer the chance to discover many novel bioconversions, concluding stereo/regio-specific conversions of a range of substrates such as cyclic sugars, polyalcohols, and steroids. This is especially significant given the move toward developing “cleaner” synthetic routes based, where possible, on “white,” or more environmentally friendly biotechnologies [28,36,37]. Timeline compressions in the development cycle of pharmaceuticals, in combination with a missing broad strain and enzyme choice, result in the fact that conversions typically represent the second generation process choice in the manufacturing of small molecule pharmaceuticals. As the enzyme is the first and foremost functional element in biotransformation for small molecules, novel biocatalysts especially oxidoreductases and lyases, are needed (other enzymes that have been exploited for organic synthesis, as well as the type of reaction catalyzed, are summarized in Table 5.2). Good examples of the replacement of traditional organic processes by a greener biocatalytic alternative include the thermolysin-­catalyzed synthesis of the low calorie sweetener aspartame, the production of acrylamide and nicotinamide (assisted by nitrile hydratases), the synthesis of the noncariogenic sweetener isomaltulose by sucrose mutases, the production of biopolymers such as polylactic acid, the synthesis of semisynthetic penicillins and cephalosporins, the transformation of natural and synthetic fibers, the pulp kraft-bleaching and recycling of paper, and the multistep synthesis of polyketide and glycopeptide antibiotics [29,37–39]. To overcome the problem of competing chemical reactions, the traditional frontiers for enzymatic conversions have been protein activation and stabilization, and reaction specificity [40]. For example, using protein engineering and high-throughput screening it is currently possible to create stereoselective enzymes with broadly extended substrate specificities, which are useful in organic synthesis, stable hydroxynitrile hydrates for the synthesis of, e.g., substituted R-mandelic acids, and hydrolases capable of enantioselective carbon–carbon bond formation or selective oxidation processes [27,41]. Moreover, in synthetic reactions, involving either oxidoreductases, expensive redox cofactors, or where the whole cells are needed, biotechnological developments—from upstream (strain, cell, and organism development) and midstream (fermentation and other unit operations) to downstream processes— will significantly benefit from the application of white biotechnology in chemical transformations [42]. Additionally, combinatorial immobilization techniques are providing effective methods for optimizing operational performance, in addition to the aiding the recovery and re-use of biocatalysts [40]. It should also be taken into consideration that as the drug targets are so variable and complex, industrial emphasis have been directed to the development and exploration of new domino and multicomponent reactions (MCRs) for the biosynthesis of medically important unusual antibiotics. Therefore, new catalytic synthetic methods in organic chemistry that satisfy increasingly stringent environmental constraints are in great demand by the pharmaceutical and chemical industries [38,43,44]. In addition, novel catalytic procedures are necessary to produce the emerging classes of compounds that are becoming the targets of molecular and biomedical research. Independently of the application, an ideal biocatalyst requires a high specific activity and stability and minimal substrate and product inhibition, and, very importantly, in terms of synthetic utility, a high stereospecificity. Although many enzymes have now been proven to be useful for the synthesis, one still cannot use enzymes for the formation of every desired linkage or for the resolution of any racemic mixture. Moreover, although many enzymes have been well characterized with regard to substrate specificity and stereoselectivity, they may still exhibit some unpredictable

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features with unnatural substrates. As we will discuss later, the success of metagenomics in finding new enzymatic activities may play a key role for the generation of new products and processes that were until recently hidden from us [20].

5.4 Main Chemical Conversions Mediated by Enzymes: Putative “Rare” Conversions It is likely that ongoing enzyme discovery will further enhance the catabolic potential of biotechnology toward synthetic conversions and ultimately contribute to the sustainable and environmentally friendly practices. Below we discuss the most important enzymes and conversions on which the future research will be focused.

5.4.1  Hydrolytic Enzymes in Enantio-Transformations Introduction of chirality and/or synthesis-resolution of enantiomerically pure intermediates and products are of practical importance for pharmaceuticals, agrochemicals, and other industries (Figure 5.3a). Of particular interest are esterases and lipases, which are considered to be a third important group of industrial enzymes suitable for commercial exploitation. Both types of enzymes belong to the family of carboxylic ester hydrolases (EC 3.1.1.), which catalyze the cleavage of ester bonds of lipids and other organic compounds, and are divided into 79 enzymatic families according to the specific bond, moiety, and substrate they hydrolyze [45]. Although their primary natural function is the hydrolysis of acylglycerols, the main interest in their application stems from their high activity and stability in nonaqueous ­systems, which allow important biotransformations such as preparation of enantiopure (a)

Enantiomeric resolution mediated by esterases and lipases OH

OH

OH

O

O

(b)

Synthesis of oligosaccharides by glycosidases, transglycosidases, and glycosyltransferases OH O HO OH OH HO O GluFa O OH O OH HO O OH O OH O Glucosa HO HO OH OH 30 Glucosa 25 Maltosa 20 Maltotriosa 15 mV

HO HO

10 5 0 0

5 10 15 20 25 Tiempo de retención (min)

OCH3

OH

OCH3

30

+

O

(c)

OH O

+ CH3OH

Regiospecific hydroxylation mediated by mono-oxygenases COOH

COOH

COOH COOH NH2

COOH NH2 N

OH OH OH

OH

OH

Enantiospecific reduction of carboxylic groups to chiral alcohols by alcohol dehydrogenases OH O F O O O F F F OH O O F F F O F F

Figure 5.3  A few examples of preparative and industrial applications of enzymes. (a) Esterase-lipase mediated resolution of chiral synthons. (b) Enzymatic synthesis of glycosidic-formation enzymes. (c) Regiospecific hydroxylation and enantiospecific reduction by mono-oxygenases and alcohol dehydrogenases.

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compounds from racemic pairs, prochiral (precursors to chiral) or meso compounds, or diastereomeric mixtures [32,33]. Hydrolytic enzymes also effectively catalyze enantiocomplementary reverse hydrolysis (esterification, transesterification, aminolysis, or amidation), providing access to both enantiomers of a desired product [46]. Moreover, in order to produce enantiopure-chiral amines and alcohols as well as phospho-, glyco-, and lipopeptide conjugates, lipase/esterase-catalyzed enantioselective reactions for temporary protection-deprotection of alcohol and amines (using appropriate protecting groups) have recently been reported [47–49]. Further, the regiospecific esterification of carbohydrates by the enzymecatalyzed transesterification of sugars, may be obtained with an appropriate selection of lipases [46]. Besides esterases and lipases, two other important hydrolytic enzymes are proteases and penicillin acylases. Proteases keep their status as important catalysts for peptide synthesis in thermodynamically controlled processes or kinetically controlled aminolysis; similarly to other hydrolases such as esterases and lipases, proteases not only serve as hydrolytic enzymes, hydrolyzing peptide bonds in vivo, but also facilitate amide- or ester-bond-forming reactions under specific conditions in vitro, being particularly interesting for the formation of unnatural amino acids through the peptide bond formation. Moreover, apart from lipases and carboxylesterases, serine proteases, especially those of the subtilisin-family, have also been successfully employed in sugar acylation processes, even with long-chain fatty acids [46]. The last example of hydrolytic enzymes applied in synthetic industrial processes are penicillin acylases. Although, they are broadly appreciated for the synthesis of medically important antibiotics through β-lactam cleavage, they harbor a few practical limitations, such as the narrow pH profile (they are active within a narrow window of pH) and that the resolution of antibiotics is kinetically controlled yielding max. 50% yield [50]. Although in silico analysis, modeling techniques, protein engineering, and medium development can aid the “correct” stereochemical course of the reaction along with the improvement of thermal and/or solvent stability and facilitate the insight into potential substrate or enzyme modifications that may increase selectivity [4], these are less attractive than the discovery of new enzymes containing unnatural amino acids or functionalities that yield new structures and cannot be obtained through existing technologies.

5.4.2 Rare Conversions Using Carbohydrate Modifying Enzymes: Production of Rare Carbohydrates It is known that a common oligosaccharide is a carbohydrate consisting of two to ten monosaccharide residues linked by O-glycosidic bonds [51,52]. The development of efficient and scalable synthetic processes for addressing oligosaccharides in the food (prebiotics, sweeteners, stabilizers, bulking agents, etc.) and pharmaceuticals (e.g., therapeutics in prevention of infection, neutralization of toxins, and for immunotherapies) is of great interest [53]. Considering the structural diversity of oligosaccharides (three different amino acids would allow the synthesis of only six different peptides, whereas three different hexopyranose moieties would yield up to 720 trisaccharides), the stereo- and regioselectivity of enzymes is considered a valuable alternative to chemical synthesis—which needs complex protection and deprotection steps—for the preparation of structurally well-defined oligosaccharides (Figure 5.3b). Actually, enzymatic processes are the preferred choice in the food industry for the production of most important oligosaccharides. Biotransformation of carbohydrates is a classical example of the application of the regiospecificity of enzymes [54]. In vivo synthesis of glycosidic bonds is performed by glycosyltransferases (EC 2.4.) [55,56]. These enzymes catalyze the coupling (by a transfer reaction) of a glycosyl donor to an acceptor molecule forming a new glycosidic bond with regio- and stereo-selectivity. According to the nature of the sugar residue being transferred, glycosyltransferases are divided into hexosyltransferases (EC 2.4.1.), pentosyltransferases (EC 2.4.2.), and those transferring other glycosyl groups (EC 2.4.99.). Depending on the nature of the donor molecule, glycosyltransferases are classified into three main mechanistic groups: (1) Leloirtype glycosyltransferases, which require sugar nucleotides (e.g., UDP-glucosyltransferase); (2) non-­Leloir glycosyltransferases, which use sugar-1-phosphates (e.g., phosphorylases); and (3)  transglycosidases,

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which employ nonactivated oligosaccharides (e.g., sucrose, starch, etc.) as glycosyl donors. A distinctive feature of transglycosidases, compared with Leloir and non-Leloir glycosyltranferases, is that they also display some hydrolytic activity, which can be regarded as a transfer of a glycosyl group from the donor to water. It is noteworthy to mention that in terms of reaction mechanism transglycosidases belong to the same group that glycosidases (EC.2.), a group of hydrolases that catalyze with exquisite sterereoselectivity the hydrolysis of glycosidic bonds in oligo- and polysaccharides. According to classification Henrissat based on amino acid sequence comparisons transglycosidases and glycosidases constitute the “glycoside hydrolase family (GH family),” with more than 2,500 enzymes [57]. In vitro oligosaccharide synthesis can be performed with glycosyltransferases and glycosidases. There are several problems associated with the use of glycosyltransferases of Leloir and non-Leloir type: (1) the requirement of sugar nucleotides or sugar phosphates as substrates, whose synthesis is rather difficult and expensive; and (2) the inhibitory effect of the nucleotide phosphate released; (3) the limited availability of these enzymes [58]. Glycosidases are widely employed for oligosaccharide synthesis, as under appropriate conditions the normal hydrolytic reaction can be reversed toward glycosidic bond synthesis [54]. This can be achieved by thermodynamic control (using low-water concentrations) or by kinetic control (using activated glycosyl donors at high concentrations). Despite the broad specificity of glycosidases and their availability, the application of these catalysts is often limited by low yields and poor regioselectivity. 5.4.2.1  Production of Rare Monosaccharides Along with the production of other sugars, the enzymatic transformations of monosaccharides have become an important bioprocess. In the past few years, the medical applications of L-carbohydrates and derived nucleosides have greatly increased [59]. Several nonmodified and modified L-sugars, e.g., L-sorbose, L-arabinose, 2-deoxy-2-fluoro-5-methyl-β-L-arabinofuranosyl uracil (L-FMAU), and L-arabinose condensed with cyanohydrin and nitromethane have been shown to be potent antiviral agents and also useful in antigenic therapies and have been used as precursors for the facile synthesis of the potent glycosidase inhibitor 1-deoxygalactonojirimycin, as anti-hepatitis B virus, anti-human immunodeficiency virus (HIV), anti-HBV (hepatitis B virus) and anti-EBV (Epstein-Barr virus), and antitumor agents [59]. Rare sugars are usually as sweet as the natural sugars, but unlike them, rare sugars are either not metabolized by the body or metabolized to a lesser extent than natural sugars. Due to these features, rare sugars are desirable as low-calorie sweeteners and are well tolerated by diabetics. It was also found that L-monosaccharides have antineoplastic characteristics useful in combination with all major forms of cancer therapy including surgery, biological, chemical and radiation therapies, and hyperthermia. Other advantage of rare sugars is the absence of an objectionable aftertaste, commonly experienced with artificial sweeteners such as saccharin or cyclamates. However, in spite of the demand for these rare sugars, their commercial availability, application, or usefulness is negligible as they are expensive to prepare, and unavailable in nature. Apart from their intrinsic applications, one of the key challenges in the last decades involves the asymmetric synthesis of highly functionalized monosaccharides that bear several centers of chirality [60]. Subsequently, these sugars (i.e., monosaccharides) need to be connected through glycosidic linkages to carboxydrates or other biomolecules, such as lipids, proteins, or metabolites [49]. However, the selective formation and hydrolysis of glycosidic linkages is chemically difficult, but is a common reaction in biological systems—it is estimated that 1–2% of an organism’s genes are dedicated to glycoside hydrolases and glycosyltransferases. The application of biocatalysis for carbohydrates is, therefore, a rich field that has now been adopted to various degrees, by many researchers who are involved in carbohydrate synthesis. The key enzymes for the synthesis of monosaccharides are readily available aldolases which catalyze the selective formation of the central C–C bonds, e.g., galactose oxidase, rhamnulose-1-phosphate aldolase, sialyltransferase. These enzymes have successfully been applied to the synthesis of a variety of mono- and sialooligosaccharides [34,61]. The formation of C–C bonds by aldolases for the production of monossacharides with complete stereochemical control is of obvious importance in organic synthesis, and enzyme-catalyzed aldol addition reactions (typically between an aldehyde and a ketone) have made important contributions in this regard [62]. In aldolase-catalyzed

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reactions, the enzyme generally controls configuration of newly formed stereogenic centres. Aldolases are highly specific for the donor substrate (that is, the nucleophilic enolate) but relatively flexible with respect to the acceptor (electrophilic) group. Exploiting this enzymatic characteristic, a judicious choice of acceptor substrate and aldolase, has led to the preparation of numerous carbohydrates and mimics thereof. These molecules have further served as intermediates in the synthesis of complex bioactive molecules, such as glycosyltransferase and glycosidase inhibitors. Characteristic synthetic examples are the use of the fructose diphosphate (FDP) aldolase for the construction of cyclic imine sugars and the acetaldehyde-dependent aldolase 2-deoxyribose-5-phosphate aldolase (DERA). The latter is the only known aldolase which catalyzes condensation between two aldehydes, and which has been used in the synthesis of epothilones, a new class of anti-cancer agents of interest in the pharmaceutical industry [34]. Other synthetically useful enzymes catalyzing C–C bond formation include transaldolases, transketolases, cyanohydrin synthetases (also called oxynitrilase), and enzymes for acyloin condensation, acyltransfer, isoprenoid, and steroid assembly, β-replacement of amino acids, and many B12-dependent reactions which will not be discussed here. Apart from aldolases for the production of monosaccharides, other enzymes have been successfully applied for the formation of glycosidic linkages, namely for the production of complex sugars with prebiotic and bioactive properties (see below). 5.4.2.2  Rare Oligossacharides—Prebiotics and Bioactives Transglycosidases are the ideal biocatalysts for oligosaccharide synthesis in vitro, since they do not require special activated substrates, as they directly employ the free energy of cleavage of disaccharides (e.g., sucrose) or polysaccharides (e.g., starch) [63]. For this reason, they are the most convenient enzymes for the production of prebiotic oligosaccharides as food ingredients that are potentially beneficial to the human or animal health (prebiotics escape enzymatic digestion in the upper gastrointestinal tract and enter the colon without change to their structure) [40]. Transglycosidases exhibit the same mechanism as retaining glycosidases, resulting in net retention of anomeric configuration. The active site contains two carboxylic acid residues, located at approx. 5.5 Å apart: one acting as a nucleophile and the other as an acid/base catalyst. The reaction proceeds by a double-displacement mechanism in which a covalent glycosyl-enzyme intermediate is formed by the attack of the deprotonated carboxylate to the anomeric centre of the carbohydrate with concomitant C-O breaking of the scissile glycosidic bond [56,60]. This step is assisted by the carboxylic residue acting as a general acid. The second step is the attack of a nucleophile to the glycosyl-enzyme intermediate, which is assisted by the conjugate base of the second carboxyl residue. The nucleophilic H2O and the acceptor (normally a carbohydrate) compete for the glycosyl-enzyme intermediate. When the nucleophile is H2O, the enzyme acts as a hydrolase; when the sugar is the nucleophile, the enzyme acts as a transferase. The transferase/hydrolase ratio depends on two main factors: (1) the concentration of acceptor (high concentrations must be used to enhance glycosyl transfer), and (2) the intrinsic enzyme properties. A transglycosidase will be considered efficient if it possesses significant ability to bind the acceptor and to exclude H2O. Along with transglycosidases, the glycosyltransferases are the biocatalysts responsible for the formation of many of the glycosyl bonds in nature. However, these enzymes use either sugar nucleotides or glycosyl phosphates as their activated sugar donors and are often highly selective for the second substrate, the glycosyl acceptor. Although we have described above the limitation of those in biocatalysis, they have been demonstrated as useful biocatalysts, e.g., for the glycosylation of natural products, such as the glycopeptide antibiotics vancomycin and teicoplanin, or the angucyclic antitumor drugs urdamycin A and B by using glycorandomization strategies [60,64]. However, these enzymes in some cases require new donor substrates (glycosyl fluorides, oxazolines, and 6-oxo-glycosides) for synthetic transglycosylation reactions. Encouragingly, some of the glycosyltransferases that are involved in the biosynthesis of above natural products have been purified and have demonstrated different degrees of enzymatic promiscuity toward their substrates. Particularly important targets for glycosylation are glycoproteins with defined and homogeneous glycosylation patterns. Biocatalytic approaches have involved the enzymatic glycosylation of neoglycoproteins after site-selective chemical glycosylation

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or re-engineering of the glycosylation machinery of Pichia pastoris to produce human-like N-linked glycosylation on proteins. As a consequence of the progress in the understanding of the structures and catalytic mechanisms involved in the enzymatic synthesis of glycosidic bonds, a group of novel, directed-mutated glycosidases called glycosynthases was developed [65]. The glycosynthase concept was established in 1998 by the Whiters’ group using an exo-glucosidase as a model [66], and was further extended to endo-glycosidases by the Planas’ group [67]. A glycosynthase is a specifically mutated retaining glycosidase in which substitution of the catalytic carboxyl nucleophile by a noncatalytic residue (Ala, Gly, or Ser) renders a hydrolytically inactive enzyme, still able to catalyze the transglycosylation of activated glycosyl fluoride donors (having the opposite anomeric configuration to that of normal substrates of the parental wildtype enzyme). The yield obtained with glycosynthases reaches in some cases 95–98% [58]. The impressive amount of ­glycosidases available clearly indicates that the potential biodiversity of glycosynthases is still largely unexplored, and new applications of these enzymes will emerge in the near future [68]. Among the most challenging oligosaccharide targets for synthesis are those of the proteoglycans, such as heparin and heparan sulfate. These are negatively charged, linear polysaccharides that interact with a variety of proteins at the cell surface. An important target has been the antithrombin III binding pentasaccharide, which promotes inhibition of blood coagulation, and hence, has important therapeutic applications. The chemical synthesis of the pentasaccharide has been reported in 60 steps, with less than 0.5% overall yield [34]. The challenge for rare conversion of carbohydrates is to develop a robust screening or selection systems, and in this context, a recent report by Mayer et al. [69] describes the development of a novel agar plate-based coupled-enzyme screen to select natural glycosynthases from a library of mutants. Such developments may open the window to find wild-type enzymes derived from environmental DNA which show high ratio synthesis versus hydrolysis of glycosidic bonds.

5.4.3 Oxidative Enzymes for the Conversion of Organic Pollutants in Industrial Effluents to Useful Products Research involving enzymes such as peroxidases, laccases, and polyphenol oxidases from plants, fungi, and bacteria, aims at the development of bioremediation systems, and in addition, utilizing the waste organics found in industrial wastewaters, converting them to economically valuable products [37]. At present, wastes from the wine, olive, and petrochemical industries are used as sources of raw materials for the bioconversions. Potential products include antioxidants suitable for use as food additives or nutraceuticals. A number of oxidase enzymes are being investigated for their ability to act as biocatalysts in the hydroxylation and oxidation of aromatics to produce compounds of economic importance, particularly for production of compounds of value in the pharmaceutical and food industry (Figure 5.3c), namely ferulic acid and certain flavonoids and steroids.

5.4.4 Rare Conversions Using Fatty Acids: Production of Unique Complex Lipids and Essential Fatty Acids It is now clear that the identification of novel biochemical conversions for carboxylic acid activation may reveal new potential drug and human health targets. More generally, the identification of novel classes of enzymes and their role in the biosynthesis of complex hybrid metabolites may also reveal unknown mechanisms with which we should be able to generate metabolite diversity: fatty acid synthases (Fad) and poliketide synthases (PKSs) in combination have been demonstrated as powerful catalysts to generate hybrid metabolites, although it is still not clear how the covalent sequestered biosynthetic intermediates are transferred from one enzymatic complex to another [37,70,71]. Long-chain polyunsaturated fatty acids (PUFAs) are vital for human health [72]. Although, until now, fish and other marine oils have been the only source of PUFAs, new developments have been achieved for the biosynthesis of ω-3 and

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ω-6 families of PUFAs. Synthesis of these essential fatty acids requires the introduction of several genes responsible for the conversion of linoleic acid into arachidonic acid, or α-linolenic acid into eicosapentaenoic acid and docosahexaenoic acid. Although the three major steps for the synthesis of PUFAs are all well studied and involve sequentially a ∆6-desaturation, a chain elongation and a ∆5-desaturation by the actuation of fatty acid synthases, PKSs, and desaturases, it still remains unclear how efficient other enzymes could be for carboxylic acid activation [72].

5.4.5 Rare Conversions Using Aliphatic Groups: Incorporation of Chlorine, Fluorine, Bromide, and Iodine to Peptides-Polyketides Enzymatic incorporation of chlorine, bromide, or iodine atoms occurs during the biosynthesis of more than several thousands natural products [73]. This particular halogenation may have significant consequences for the bioactivity of these products so there is a great interest in understanding the biological catalysts that perform these reactions. With the exception of the well characterized haloperoxidases [74], most of the biosynthetic enzymes and mechanisms responsible for the halogenations have remained elusive. The crystal structures of two functionally diverse halogenases have been recently solved, providing us with new and exciting mechanistic detail [75,76]. This new insight has the potential to be used both in the development of biomimetic halogenation catalysts and in engineering halogenases, and related enzymes, to halogenate new substrates. Interestingly, these new structures also illustrate how the ­evolution of these enzymes mirrors that of the monooxygenases, where the cofactor is selected for its ability to generate a powerful oxygenating species [74]. Since their discovery, halogenated metabolites have been somewhat of a biological peculiarity and it is only now that we are beginning to realize the full extent of their medicinal value. However, only few studies report on enzymes that halogenated unactivated aliphatic groups from peptides and polyketides. A good example of an efficient enzyme catalysing rare conversions such as iodination, bromidation, and chlorination of a wide range of substrates are chloroperoxidases (CPO) [74]. Besides H 2O2dependent halogenation reactions, the enzyme catalyzes dehydrogenation reactions. These reactions can be performed in a stereoselective manner by CPO, which may also act as a catalase, facilitating the decomposition of hydrogen peroxide to oxygen and water. Furthermore, CPO catalyzes P450like oxygen insertion reactions, e.g., enantioselective epoxidation/sulfoxidation of alkanes. Although CPO have extensively been investigated during the last four decades, it should be crucial for the conversion of nonactivated compounds such as alkanes to find novel CPO-like enzymes that help to gain a better understanding of the catalytic mechanism of this class of enzymes. This is of practical importance as we know that with the exception of the well-characterized haloperoxidase, most of the biosynthetic enzymes and mechanisms responsible for the halogenation have remained elusive [74,76]. Alternatively, the isolation of novel halogenases may open new perspectives to realize the full extent of their medical value [73]. This new insight has the potential to be used for the development of biomimetic halogenation catalysts and in engineering halogenases, and related enzymes, to halogenate new substrates. We are convinced that natural evolution can efficiently select particular structures and properties to enhance or modify enzyme catalysis. Although fluorine in the form of fluoride minerals is the most abundant halogen in the Earth’s crust, only 12 naturally occurring organofluorine compounds have so far been found, and how these are biosynthesized remains a mystery [1]. The rarity of natural fluorinated products contrasts with the identification of about 3,500 naturally occurring halogenated compounds. The available fluoride is largely insoluble—e.g., sea water contains 1.3 ppm. fluoride and 19,000 ppm. chloride, which may help to explain why fluorine’s biochemistry has hardly evolved. The toxin fluoroacetate is the most ubiquitious of the small class of organofluorine compounds and has been identified in more than 40 plant species from all of the continents apart from Antarctica, but its biosynthetic fluorination pathway has not been clearly defined. Fluoroacetate is also produced by the bacterium Streptomyces cattleya when it is grown in culture medium supplemented with fluoride ions. In a recent study, an enzymatic reaction has been

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described that occurs in the bacterium S. cattleya and which catalyzes the conversion of fluoride ion and S-adenosylmethionine (SAM) to 5′-fluoro-5′-deoxyfluoroadenosine (5′-FDA). This study has for the first time revealed a fluorinase enzyme, this discovery opened up new biotechnological opportunities for the preparation of organofluorine compounds and showed the usefulness of continuation of discovery of new halogenases for both degradation and synthetic purposes [75].

5.4.6 Rare Conversions Using Amino Acids: Production of Unnatural Amino Acids The products of such biocatalytic systems have significant value as precursors of antibiotics and other high-value pharmaceuticals, particularly due to the stereospecificity of the enzymes involved and hence the chirality of the products. In this way, enantioselective amino acid syntheses using biocatalysis methods, specifically involving proteases and hydantoin-hydrolysing enzyme systems are broadly appreciated [77,78]. Unnatural amino acids are a growing class of intermediates required for pharmaceuticals, agrochemicals, and other industrial products [79]. However, no single enzyme was proven to be efficient enough to prepare these compounds broadly at scale. To address this need we need to isolate new enzymes to prepare enantiomerically pure L- and D-amino acids in high yield by deracemization of racemic starting materials (enantiomeric excess and product yields over 99%).

5.4.7  Rare Conversion with Flavonoids and Steroids: Drug Synthesis Flavonoids are among the most ubiquitous phenolic compounds found in nature [80]. These compounds have diverse physiological and pharmacological activities such as estrogenic, antitumor, antimicrobial, antiallergic, and anti-inflammatory effects. They are well-known antioxidants and metal ion-chelators. As integral constituent of the diet, flavonoids may exert a wide range of beneficial effects on human health, including protection against cardiovascular diseases and certain forms of cancer. Recent studies have shown diverse physiological and pharmacological activities of these natural compounds such as strogenic, antilipoperoxidant, antitumor, antiplatelet, antiviral, antifungal, antibacterial, antihemolytic, anti-ischemic, antiallergic, and anti-inflamatory. The biological effect is mediated by their free radical-scavenging antioxidative activities and metal ion-chelating abilities. These properties may facilitate inhibitions of certain enzymes such as lipooxygenases, cyclooxygenases, monooxygenases, xantine oxidases, aldose reductase, etc. It is therefore important to increase or decrease the biological activities of those flavonoids by enzymatic transformations. Biotransformations of numerous flavonoids catalyzed mainly by microbes and few plant enzymes have been described in four different flavonoid classes, chalcones, isoflavones, catechins, and flavones. These biotransformations represent a variety of reactions including condensation, cyclization, hydroxylation, dehydroxylation, alkylation, O-dealkylation, halogenation, reduction, deglycosidation, methylation, dehydrogenation, double-bond reduction, carbonyl reduction, glycosylation, sulfation, dimerization, or different types of ring degradations, conjugations, and reductions [80]. Future advances should be made in the discovering of enzymes able to efficiently separate racemic flavones and/or introduce chiral centers or functional groups, to achieve efficient derivatives that may be used as valuable synthons: benzaldehyde lyase (BAL, EC 4.1.2.38), biphenyl dioxygenase, dihydrodiol dehydrogenase, phenylalanine/tyrosine ammonia lyase, 4-coumrate:coenzyme A (CoA) ligase, chalcone synthase, cinnamte hydroxylase, chalcone isomerase, methyltransferase, biphenyl-2,3-dihydrodiol-2,3-dehydrogenase, O-demethylase, ­t yrosinase, horseradish peroxidase [80]. Other important derivatives, steroids, are also commonly used to treat ailments such as arthritis, skin disorders, and adrenal insufficiency. These therapeutic