Inositol Phosphates: Linking Agriculture and the Environment

Inositol Phosphates

LINKING AGRICULTURE

AND THE

ENVIRONMENT

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Inositol Phosphates LINKING AGRICULTURE

AND THE

ENVIRONMENT

Edited by

Benjamin L. Turner Smithsonian Tropical Research Institute Balboa, Ancón, Republic of Panama

Alan E. Richardson CSIRO Plant Industry Canberra, Australia and

Edward J. Mullaney United States Department of Agriculture New Orleans, USA

CABI is a trading name of CAB International CABI Head Office Nosworthy Way Wallingford Oxfordshire OX10 8DE UK Tel: +44 (0)1491 832111 Fax: +44 (0)1491 833508 E-mail: [email protected] Website: www.cabi.org

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©CAB International 2007. All rights reserved. No part of this publication may be reproduced in any form or by any means, electronically, mechanically, by photocopying, recording or otherwise, without the prior permission of the copyright owners. A catalogue record for this book is available from the British Library, London, UK. A catalogue record for this book is available from the Library of Congress, Washington, DC. ISBN-10: 1 84593 152 1 ISBN-13: 978 1 84593 152 1

Typeset by SPi, Pondicherry, India. Printed and bound in the UK by Biddles Ltd, King’s Lynn.

Contents

Contributors Preface Acknowledgements 1.

2.

3.

4.

vii ix xi

Nomenclature and Terminology of Inositol Phosphates: Clarification and a Glossary of Terms Stephen B. Shears and Benjamin L. Turner

1

Identification of Inositol Phosphates by Nuclear Magnetic Resonance Spectroscopy: Unravelling Structural Diversity Pushpalatha P.N. Murthy

7

High-performance Chromatographic Separations of Inositol Phosphates and Their Detection by Mass Spectrometry William T. Cooper, Matthew Heerboth and Vincent J.M. Salters

23

Origins and Biochemical Transformations of Inositol Stereoisomers and Their Phosphorylated Derivatives in Soil Michael F. L’Annunziata

41

5.

Isolation and Assessment of Microorganisms That Utilize Phytate Jane E. Hill and Alan E. Richardson

6.

Phytate-degrading Enzymes: Regulation of Synthesis in Microorganisms and Plants Ralf Greiner

61

78

7.

Phytases: Attributes, Catalytic Mechanisms and Applications Edward J. Mullaney and Abul H.J. Ullah

97

8.

Seed Phosphorus and the Development of Low-phytate Crops Victor Raboy

111

9.

Phytase and Inositol Phosphates in Animal Nutrition: Dietary Manipulation and Phosphorus Excretion by Animals Xin Gen Lei and Jesus M. Porres

133

v

vi

Contents

10.

Environmental Implications of Inositol Phosphates in Animal Manures April B. Leytem and Rory O. Maguire

150

11. Ligand Effects on Inositol Phosphate Solubility and Bioavailability in Animal Manures Thanh H. Dao

169

12. Inositol Phosphates in Soil: Amounts, Forms and Significance of the Phosphorylated Inositol Stereoisomers Benjamin L. Turner

186

13.

207

Abiotic Reactions of Inositol Phosphates in Soil Luisella Celi and Elisabetta Barberis

14. Interactions Between Phytases and Soil Constituents: Implications for the Hydrolysis of Inositol Phosphates Timothy S. George, Hervé Quiquampoix, Richard J. Simpson and Alan. E. Richardson

221

15.

Plant Utilization of Inositol Phosphates Alan E. Richardson, Timothy S. George, Iver Jakobsen and Richard J. Simpson

242

16.

Inositol Phosphates in Aquatic Systems Ian D. McKelvie

261

Index

279

Contributors

Barberis, Elisabetta, University of Turin, DIVAPRA Chimica Agraria, via Leonardo da Vinci 44, Grugliasco, 10095 Torino, Italy Celi, Luisella, University of Turin, DIVAPRA Chimica Agraria, via Leonardo da Vinci 44, Grugliasco, 10095 Torino, Italy Cooper, William T., Department of Chemistry and Biochemistry, Florida State University, Tallahassee, FL 32306, USA Dao, Thanh H., United States Department of Agriculture–Agricultural Research Service, Beltsville Agricultural Research Center, Room 121, 10300 Baltimore Avenue, Building 306 BARC-EAST, Beltsville, MD 20705, USA George, Timothy S., Scottish Crop Research Institute, Invergowrie, Dundee DD2 5DA, UK Greiner, Ralf, Federal Research Centre for Nutrition and Food, Centre for Molecular Biology, Haid-und-NeuStraße 9, D 76131 Karlsruhe, Germany Heerboth, Matthew, Department of Chemistry and Biochemistry, Florida State University, Tallahassee, FL 32306, USA Hill, Jane E., Environmental Engineering Program, Yale University, 9 Hillhouse Avenue, PO Box 8286, New Haven, CT 06520, USA Jakobsen, Iver, Risø National Laboratory, Biosystems Department, Roskilde, DK 4000, Denmark L’Annunziata, Michael F., The Montague Group, PO Box 5033, Oceanside, CA 92052, USA Lei, Xin Gen, Department of Animal Science, Morrison Hall 252, Cornell University, Ithaca, NY 14853, USA Leytem, April B., United States Department of Agriculture–Agricultural Research Service, Northwest Irrigation and Soils Research Laboratory, 3793 N. 3600 E., Kimberly, ID 83341, USA Maguire, Rory O., Crop and Soil Environmental Sciences, Virginia Tech, Box 0404, Blacksburg, VA 24061, USA McKelvie, Ian D., Water Studies Centre and Chemistry Department, School of Chemistry, Monash University, Clayton, Victoria 3800, Australia

vii

viii

Contributors

Mullaney, Edward J., United States Department of Agriculture–Agricultural Research Service, Southern Regional Research Center, 1100 Robert E. Lee Blvd, New Orleans, LA 70124, USA Murthy, Pushpalatha P.N., Department of Chemistry, Michigan Technological University, 1400 Townsend Drive, Houghton, MI 49931, USA Porres, Jesus M., Departamento de Fisiología, Universidad de Granada, Granada, Spain Quiquampoix, Hervé, Unité de Science du Sol, INRA-ENSAM, 2 Place Pierre Viala, 34060 Montpellier Cedex 1, France Raboy, Victor, United States Department of Agriculture–Agricultural Research Service, Small Grains and Potato Germplasm Research Unit, 1691 S. 2700 W., Aberdeen, ID 83210, USA Richardson, Alan E., CSIRO Plant Industry, PO Box 1600, Canberra, ACT 2601, Australia Salters, Vincent J.M., National High Magnetic Field Laboratory and Department of Geological Sciences, Florida State University, Tallahassee, FL 32306, USA Shears, Stephen B., Laboratory of Signal Transduction, National Institute of Environmental Health Sciences, NIH, DHSS, Research Triangle Park, PO Box 12233, NC 27709, USA Simpson, Richard J., CSIRO Plant Industry, PO Box 1600, Canberra, ACT 2601, Australia Turner, Benjamin L., Smithsonian Tropical Research Institute, Apartado 0843-03092, Balboa, Ancón, Republic of Panama Ullah, Abul H.J., United States Department of Agriculture–Agricultural Research Service, Southern Regional Research Center, 1100 Robert E. Lee Blvd, New Orleans, LA 70124, USA

Preface

Inositol phosphates are a group of organic phosphorus compounds found widely in the natural environment. They are common in eukaryotic organisms, especially plants, where they constitute most of the phosphorus in seeds. Soils and aquatic sediments also contain large amounts of inositol phosphates, some of which occur in forms that have not been detected anywhere else in nature. The abundance of inositol phosphates in nature means that they are of widespread interest in the ecological and environmental sciences. However, it is in the science of animal nutrition that inositol phosphates have become a topic of considerable interest. This stems from the fact that monogastric animals cannot digest phytate (salts of myo-inositol hexakisphosphate), the most abundant inositol phosphate in cereal grains. Supplemental phosphate is therefore required in the diets of pigs and poultry to maintain productivity. A consequence of phosphate supplementation is that animal manure can contain considerable concentrations of phosphorus. Not only does this represent a financial loss to the producer, but it also contributes to one of the most pervasive forms of environmental pollution from modern agriculture. Long-term application of manure to agricultural land leads to an accumulation of phosphorus in the soil and a gradual increase in phosphorus transport in runoff to water bodies. Such diffuse pollution is now widespread and there are numerous examples of regional-scale water quality deterioration in areas of intensive livestock operations. Two well-publicized examples are the Chesapeake Bay, USA, and the Gippsland Lakes, Australia. In both cases the problems have been severe and public – the high-profile detection of the neurotoxin-producing dinoflagellate Pfiesteria piscicida in the Chesapeake Bay being a particular cause for concern. To address this issue, several strategies of dietary manipulation have been developed to improve the ability of monogastric animals to digest phytate. These include the use of ‘low-phytate’ grains – mutants selected for the low concentration of inositol phosphate in their seed – and the development of transgenic animals that produce phytase, an enzyme that degrades phytate but is not naturally present in the guts of monogastric animals. By far the most successful strategy, however, has been the supplementation of animal diets with a microbial phytase. This is now standard practice in most large-scale animal feeding operations and is even mandated by law in some states of the USA. It has proved to be extremely effective in reducing phosphorus excretion in manure and has the added benefit of improving mineral nutrition by releasing metals from complexation with phytate. Despite the wealth of information on inositol phosphates in animal nutrition, the environmental impacts of manure-derived inositol phosphates and associated dietary manipulations are not well understood. In particular, the fate of the large amount of inositol phosphates being cycled through

ix

x

Preface

agriculture, especially in regions with high animal densities, is unknown. Importantly, only a handful of studies have assessed the impact of diet manipulation on phosphorus dynamics in the environment. Inositol phosphates are rapidly and strongly stabilized in soil, which means that they are often considered to be biologically unavailable and unlikely to be transported in runoff to water bodies. Yet there is growing evidence that inositol phosphates are not as recalcitrant in the environment as once thought. It is now clear that many terrestrial microorganisms, including those associated with plants, have the capacity to use inositol phosphates. This trait appears widespread, although its ecological implications await investigation. Similarly, when inositol phosphates are transported in runoff to water bodies, they can degrade rapidly and contribute to the nutrition of cyanobacteria and other aquatic organisms linked to eutrophication. Inositol phosphates can therefore no longer be considered ecologically or environmentally benign. Given the water quality problems associated with intensive livestock production and the widespread adoption of dietary modifications, there is an urgent need to improve our understanding of inositol phosphates in the environment. This was addressed at a conference held in August 2005 in Sun Valley, Idaho, USA, sponsored by the Soil Science Society of America. The meeting, entitled ‘Inositol Phosphates in the Soil–plant–animal System: Linking Agriculture and Environment’, was attended by scientists from a diverse range of disciplines with a common interest in inositol phosphates. This book is the output from that conference. Written by the invited speakers, it brings together critical reviews on the major topics in inositol phosphates in agriculture, ecology and the environment. The chapters cover three major themes: 1. State-of-the-art analytical methodology for assessing inositol phosphates in environmental samples, including nuclear magnetic resonance (NMR) spectroscopy and mass spectrometry. 2. Inositol phosphates in animal nutrition, including the latest research on plant and microbial phytases, their interactions in soil and the manipulation of animal diets with phytase supplements and low-phytate grains. 3. Inositol phosphates in the environment, including the amounts, forms and behaviour in soils and aquatic systems, their biological availability and the fate of manure-derived inositol phosphates in the environment. By covering all major aspects of inositol phosphates in agriculture and the environment, the book will serve as a unique reference source on this emerging topic. We hope that it will benefit those trying to unravel the complexity of inositol phosphates in the environment and reveal what is already known to a wider audience. The inositol phosphate conference in 2005 was held a quarter of a century after the publication of Dennis Cosgrove’s seminal text Inositol Phosphates: Their Chemistry, Biochemistry, and Physiology (Elsevier Scientific, Amsterdam). Formerly of CSIRO Plant Industry in Canberra, Australia, Cosgrove devoted his career to understanding inositol phosphates in the environment (an obituary can be found in Soil Biology and Biochemistry, vol. 14, pp. 77–78). His pioneering work in the two decades after he moved with his family from England to Australia in 1955 laid the foundations for many of the topics in this volume. His death in 1981 at the age of 56 marked the end of an era for studies on inositol phosphates in the environment, but his discoveries remain an inspiration to scientists in this field. We hope that this volume will go some way towards reinvigorating interest in these fascinating compounds. Benjamin L. Turner Smithsonian Tropical Research Institute, Balboa, Ancón, Republic of Panama Alan E. Richardson CSIRO Plant Industry, Canberra, Australia Edward J. Mullaney United States Department of Agriculture, New Orleans, USA

Acknowledgements

The meeting would not have been possible without generous support from the Soil Science Society of America through the Bouyoucos Conference fund. Bouyoucos conferences were established to facilitate an intense, highly focused examination of a topic of critical importance to soil science. Scientists with a common interest are brought together in a forum that is not typically possible at large scientific meetings, with the aim of establishing personal relationships and promoting the free exchange of ideas. We hope this latest Bouyoucos Conference fulfilled these ideals. Additional funding was provided by the Agricultural Research Service of the United States Department of Agriculture through a Professional Activities grant, and the Sun Valley Resort generously provided their conference facilities without charge. We thank those who gave their time to peer-review chapters for this volume – their expertise has contributed to the technical excellence of its contents. Finally, we thank all the delegates at the inositol phosphate conference for contributing to a vibrant and stimulating few days, and we look forward to the next meeting.

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1

Nomenclature and Terminology of Inositol Phosphates: Clarification and a Glossary of Terms Stephen B. Shears1 and Benjamin L. Turner2 1

Laboratory of Signal Transduction, National Institute of Environmental Health Sciences, NIH, DHSS, Research Triangle Park, PO Box 12233, NC 27709, USA; 2 Smithsonian Tropical Research Institute, Apartado 0843-03092, Balboa, Ancón, Republic of Panama

In a book like this one, which brings together reviews from scientists working in such diverse areas as analytical chemistry, biochemistry, agronomy and environmental science, the consolidation of terminology is of considerable importance. This chapter reviews the nomenclature of inositol phosphates and provides a glossary of the terms that are used throughout this book.

An Overview of Inositol Phosphate Nomenclature Much of what follows is based on the recommendations of the International Union of Pure and Applied Chemistry (IUPAC) (IUPAC–IUB Commission on Biochemical Nomenclature (CBN), 1973, 1977; Nomenclature Committee of the International Union of Biochemistry, 1989). Previous conferences, most notably the ‘Chilton Conference on Inositol and Phosphoinositides’, held in Dallas, Texas, USA, in 1984, have permitted the use of inositol phosphate nomenclature that is not IUPAC-approved (Agranoff et al., 1985). The audience at the Chilton conference were largely animal biochemists and the number of known inositol-containing compounds was far smaller than today. The authors may therefore have underestimated the potential for confusion

that lay ahead. It is now arguable that the Chilton meeting was a missed opportunity to enforce a much-needed, unified nomenclature. Inositol phosphate terminology continues to be misused even in the recent literature. For example, ‘phosphoinositide’ is a term that was intended to refer only to the inositol lipids (IUPAC–IUB Commission on Biochemical Nomenclature (CBN), 1977). Instead, conceptual difficulties arise when phosphoinositide is incorrectly used to describe inositol phosphates (e.g. De Camilli et al., 1996; Luttrell and Lefkowitz, 2002; Liu et al., 2004), especially as the physicochemical properties and biological actions of these soluble inositol derivatives are markedly different from those of the membrane-bound inositol lipids. Unfortunately, even such esteemed bodies as IUPAC are not immune from error; their intentions were confounded somewhat when the inositol lipid used to illustrate nomenclature was not the naturally occurring D-enantiomer, but the unnatural L-version (see Agranoff, 1978). The adoption of a consistent nomenclature clearly cannot eliminate mistakes, but it is an important first step towards limiting their frequency. The inositol phosphate literature also contains a number of examples of the misuse of chemical nomenclature, so clarification is appropriate. For example, a newcomer to the field would be

©CAB International 2007. Inositol Phosphates: Linking Agriculture and the Environment (eds B.L. Turner, A.E. Richardson and E.J. Mullaney)

1

2

S.B. Shears and B.L. Turner

forgiven for assuming that ‘IP6’ has only a single phosphate at the 6-position, when the abbreviation is incorrectly defined as ‘inositol-6-phosphate’ (Lee et al., 2005). The structural relationship between the scyllo- and myo-forms of inositol hexakisphosphate can be misunderstood when they are erroneously described as ‘conformers’ (Fisher et al., 2002). Of particular importance to this book is the term ‘phytate’. This refers to any salt of myoinositol hexakisphosphate. However, it has sometimes been considered that phytate is synonymous with phytin (Yoshida et al., 1999), even though the latter term was introduced originally to describe calcium/magnesium phytate, which was thought to comprise much of the phytate in seeds (Ashton, 1976). Further examples of the continued confusion in this field caused by incorrect terminology are given in a recent review (Michell et al., 2006). A glossary of terms is provided below as a prelude to this volume. A determined effort has been made to ensure that these terms are consistently deployed throughout the various chapters.

Glossary Conformer. This is one particular spatial arrangement of a molecule in space at any particular moment. For example, two conformers of myo-inositol are the so-called ‘chair’ and the ‘boat’ arrangements of the ring, the former being thermodynamically favourable. A switch between different conformers involves rotation around single bonds, but no chemical bonds are broken (if bonds were rearranged, and hence the configuration changed, the two molecules would be stereoisomers). For inositols, the end point of a conformational change can be a ring-flip, which involves the conversion between two alternate chair conformations. This occurs for myoinositol hexakisphosphate when solution pH increases past a critical value, whereupon the phosphates switch from being in a 5-equatorial/1-axial arrangement to a 5-axial/1-equatorial grouping (see Murthy, Chapter 2, this volume). Epimer. This is a special case of a pair of stereoisomers having two or more stereogenic centres, but differing at only one of these. For example, myo-inositol and scyllo-inositol are epimers, because of differences in the spatial

positioning of chemical bonds at one of their six stereogenic centres. Epimerization. The process by which two epimers are interconverted. Inositol. A cyclitol (cyclohexanehexol) with a hydroxyl group associated with each of the six carbon atoms on the ring. See also myo-inositol and scyllo-inositol. Inositol phosphate. The addition to the inositol ring of an ascending number of phosphate groups gives rise to a series of phosphorylated compounds (Table 1.1). The multiplicative prefixes (no part of which should be italicized) highlight the fact that each carbon atom has only one phosphate attached to it. Thus, ‘bis’, which is Latin in origin, means twice; ‘tris’ is Greek, meaning thrice or three times; and ‘kis’ is a general prefix from Greek that means times (Sarma, 2004). This distinguishes ‘n’ from ‘n-times’. Thus, if there were an inositol derivative with a chain of three phosphates attached to a single carbon atom, it would be a triphosphate, not a trisphosphate. The reader who is new to this field may be relieved to know that inositol triphosphates have not been detected (yet). However, diphosphate groups can be attached to the inositol ring (Table 1.1). These ‘inositol pyrophosphates’ occur naturally inside cells from a wide range of organisms (Shears, 2005). Throughout this book the term inositol phosphate is used in a general sense for all phosphorylated inositols present in environmental samples. myo-Inositol. This is one of the nine possible stereoisomers of cyclohexanehexol (Fig. 1.1). In the literature, when the exact nature of the stereoisomer is not defined, it can typically be presumed to be myo-inositol. In fact ‘Ins’ is an IUPAC-approved term for myo-inositol (Nomenclature Committee of the International Union of Biochemistry, 1989). The ‘Ins’ abbreviation is not used in this volume, so as not to undervalue the significance of the other stereoisomers that figure prominently in the environment. myo-Inositol hexakisphosphate. A compound in which all six hydroxyl groups of myoinositol are esterified as phosphates. myo-Inositol hexakisphosphate is a systematic name and is also popular in the cell-signalling literature (Irvine and Schell, 2001). Outside that field, this compound is more usually known as phytic acid. This is strictly defined as myo-inositol hexakis (dihydrogen phosphate), but the commonly used myo-inositol hexakisphosphate is used in this book.

Nomenclature and Terminology

3

Table 1.1. The myo-inositol phosphates and their accepted abbreviations. Number of phosphate groups

Full name myo-Inositol myo-Inositol monophosphate myo-Inositol bisphosphate myo-Inositol trisphosphate myo-Inositol tetrakisphosphate myo-Inositol pentakisphosphate myo-Inositol hexakisphosphate Diphospho-myo-inositol tetrakisphosphate Diphospho-myo-inositol pentakisphosphate Bis-diphospho-myo-inositol tetrakisphosphate

IUPAC abbreviationa

Common abbreviation

0 1 2 3 4 5 6

Ins InsP1b InsP2 InsP3 InsP4 InsP5 InsP6

Ins IP1 IP2 IP3 IP4 IP5 IP6

6

PP-InsP4

PP-IP4

7

PP-InsP5

IP7

8

[PP ] 2-InsP4

IP8

a

The italicization of the P denotes its use as an abbreviation for phosphate, rather than the chemical symbol for phosphorus. b Although it is not explicitly stated, we infer that InsP (without a numeric subscript) is actually the IUPAC-preferred abbreviation for myo-inositol monophosphate. However, we recommend InsP1, to avoid confusion with ‘InsP’, which is sometimes incorrectly used as a collective abbreviation for inositol phosphates (e.g. Tavares et al., 2002; Woodcock et al., 2003).

myo-Inositol HO

scyllo-Inositol HO

HO

HO OH

HO

neo-Inositol

OH

OH

L-chiro-(−)-Inositol

HO

HO

OH

OH

HO

OH

D-chiro-(+)-Inositol

HO

OH

epi-Inositol

HO

HO

OH

OH

HO

HO

OH HO

OH

OH HO OH

OH

HO OH

OH

HO

OH

OH

HO allo-Inositol

muco-Inositol HO

OH

HO

cis-Inositol OH

HO

HO

HO

HO HO

OH HO

OH

OH

OH HO

Fig. 1.1. The nine stereoisomeric forms of inositol.

OH

HO

OH

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S.B. Shears and B.L. Turner

Phytase. An enzyme (myo-inositol hexakisphosphate phosphohydrolase) that initiates the cleavage of one or more phosphate groups from myo-inositol hexakisphosphate. Several phytases are now known to exist and are described in detail in this volume (see Mullaney and Ullah, Chapter 7). Some authors prefer the term phytate-degrading enzyme. Phytate. This refers to any salt of phytic acid. Phytate can be soluble or insoluble and can occur in both dissolved and precipitated forms. Insoluble phytate involves polyvalent cations (e.g. iron phytate), whereas soluble phytate usually involves monovalent cations (e.g. sodium phytate). However, a recent study concluded that the neutral pentamagnesium salt is the predominant soluble form in animal cells (Torres et al., 2005). Nevertheless, phytate will precipitate out of solution, in a pH-dependent manner, once a critical concentration of divalent cations is exceeded. In most cases, myo-inositol hexakisphosphate exists as a salt, in both precipitated and dissolved forms, and can thus be termed phytate. However, to avoid confusion, the term phytate is used in this volume only when additional information about the cation or solubility is known. Phytate-degrading enzyme. This is an alternative term for phytase that is preferred by some authors when the in vivo function of the enzyme has not been unambiguously demonstrated (see Greiner, Chapter 6, this volume). Phytic acid. This is a non-systematic but widely used alternate name for the free-acid form of myo-inositol hexakisphosphate. As the salt-free form is unlikely to occur widely in nature, the term myo-inositol hexakisphosphate is preferred over phytic acid in this volume, although the term phytate is used when the salt is known. Phytic acid should not be used to describe other phosphorylated stereoisomers such as scylloinositol hexakisphosphate. Phytin. This term was originally introduced to describe ‘insoluble’ calcium/magnesium phytate deposits in the globoids of plant seed (e.g. Ashton, 1976). The term is largely obsolete, because phytate in the seeds of many species is now known to consist predominantly of magnesium/potassium salts (Ockenden et al., 2004). The use of ‘insoluble’ as an absolute description of this material also seems unwarranted, as the deposits are mobilized during seed germination. Positional isomers. This is a form of structural isomerism in which side chain groups (in this

case phosphates) are found attached to different carbons of the inositol ring. That is, atoms are bonded together in a different order, as opposed to stereoisomers, in which the connectivity is the same. myo-Inositol 1,3,4,5-tetrakisphosphate and myo-inositol 3,4,5,6-tetrakisphosphate are examples of positional isomers. The numbering of the carbon atoms follows rules developed by IUPAC (IUPAC–IUB Commission on Biochemical

2 1

6 3 5 4 (a)

HO HO 6

2

OH

1 4

HO

3

5

OH

OH (b) Fig. 1.2. (a) Agranoff’s turtle and (b) myo-inositol. (From Shears, 2004.) In its most stable chair conformation with 1-axial and 5-equatorial hydroxyl groups, myo-inositol has been said to resemble a turtle (Agranoff, 1978). International Union of Pure and Applied Chemistry (IUPAC) rules state that the 1-D-numbering of each carbon begins with the turtle’s front right flipper and proceeds in an anticlockwise direction around the ring (viewed from above). The axial hydroxyl is therefore represented by the turtle’s head (position number 2) and the equatorial hydroxyls by the limbs and tail. For further details of the numbering system and stereochemistry of the inositol phosphates the reader is referred to the IUPAC recommendations (IUPAC–IUB Commission on Biochemical Nomenclature (CBN), 1977; Nomenclature Committee of the International Union of Biochemistry, 1989) and comprehensive reviews published elsewhere (Parthasarathy and Eisenberg, 1991; Murthy, 2006).

Nomenclature and Terminology

Nomenclature (CBN), 1973; Nomenclature Committee of the International Union of Biochemistry, 1989). Agranoff’s turtle (see Agranoff, 1978; Shears, 2004) provides a timeless, visual mnemonic to the numbering of myo-inositol (Fig. 1.2). Unfortunately, there are no such aids for the other stereoisomers of inositol. scyllo-Inositol. This is one of the nine possible stereoisomers of cyclohexanehexol (inositol). It differs from myo-inositol at only one stereogenic centre – i.e. it is an epimer – and is unique in that it has a stable chair conformation in which all six hydroxyl groups are equatorial to the plane of the ring (Fig. 1.1). Stereoisomer. This refers to compounds that have the same chemical formula, the same atoms and the same connectivity, but differ in the fixed spatial positioning of bonds at a particular stereogenic carbon (for the inositols, a stereogenic carbon is one lacking a plane of symmetry). Hydroxyl groups on the inositol ring can be oriented in either an axial or equatorial manner, which gives nine possible

5

stereoisomers (Fig. 1.1). These stereoisomers are distinguished by a configurational prefix, which must be italicized (IUPAC–IUB Commission on Biochemical Nomenclature (CBN), 1973). The most abundant stereoisomer in nature is myoinositol, but several others occur in plants and animals. Only four inositol stereoisomers (myo-, neo-, scyllo- and D-chiro-) occur naturally in phosphorylated forms, predominantly in soils (see L’Annunziata, Chapter 4, and Turner, Chapter 12, this volume). Turtle. A marine reptile that provides an aide-mémoire for easy recall of the nomenclature for numbering the carbon atoms that comprise the myo-inositol ring (Fig. 1.2; Agranoff, 1978; Shears, 2004).

Acknowledgements We thank Dr Andrew Riley, University of Bath, UK, and Victor Raboy, USDA–ARS Aberdeen, USA, for their valuable contributions.

References Agranoff, B.W. (1978) Textbook errors: cyclitol confusion. Trends in Biochemical Sciences 3, N283–N285. Agranoff, B.W., Eisenberg, F. Jr, Hauser, G., Hawthorn, J.N. and Michell, R.H. (1985) Comment on abbreviations. In: Bleasdale, J.E., Eichberg, J. and Hauser, G. (eds) Inositol and Phosphoinositides: Metabolism and Regulation. Humana Press, Totowa, New Jersey, pp. xxi–xxii. Ashton, F. (1976) Mobilization of storage proteins of seeds. Annual Reviews in Plant Physiology 27, 95–117. De Camilli, P., Emr, S.D., McPherson, P.S. and Novick, P. (1996) Phosphoinositides as regulators in membrane traffic. Science 271, 1533–1539. Fisher, S.K., Novak, J.E. and Agranoff, B.W. (2002) Inositol and higher inositol phosphates in neural tissues: homeostasis, metabolism and functional significance. Journal of Neurochemistry 82, 736–754. Irvine, R.F. and Schell, M. (2001) Back in the water: the return of the inositol phosphates. Nature Reviews Molecular Cell Biology 2, 327–338. IUPAC–IUB Commission on Biochemical Nomenclature (CBN) (1973) Nomenclature of cyclitols. Recommendations 1973. Biochemical Journal 153, 23–31. IUPAC–IUB Commission on Biochemical Nomenclature (CBN) (1977) Nomenclature of phosphorus-containing compounds of biochemical importance. Recommendations 1976. Proceedings of the National Academy of Sciences of the United States of America 74, 2222–2230. Lee, H. J., Lee, S.A. and Choi, H. (2005) Dietary administration of inositol and/or inositol-6-phosphate prevents chemically induced rat hepatocarcinogenesis. Asian Pacific Journal of Cancer Prevention 6, 41–47. Liu, J.W., Anderson, S.N., Meulbroek, J.A., Hwang, S.M., Mukerji, P. and Huang, Y.S. (2004) Polyphosphoinositides suppress the adhesion of Haemophilus influenzae to pharyngeal cells. Lipids in Health and Disease 3 (online-only journal: doi:10.1186/1476-511X-3-20). Luttrell, L.M. and Lefkowitz, R. J. (2002) The role of beta-arrestins in the termination and transduction of G-protein-coupled receptor signals. Journal of Cell Science 115, 455–465. Michell, R.H., Heath, V.L., Lemmon, M.A. and Dove, S.K. (2006) Phosphatidylinositol 3,5-bisphosphate: metabolism and cellular functions. Trends in Biochemical Sciences 31, 52–63. Murthy, P.P.N. (2006) Structure and nomenclature of inositol phosphates, phosphoinositides, and glycosylphatidylinositols. In: Lahiri Majumder, A. and Biswas, B.B. (eds) Biology of Inositols and Phosphoinositides. Springer-Verlag, Berlin, pp. 1–20.

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Nomenclature Committee of the International Union of Biochemistry (1989) Numbering of atoms in myo-inositol. Recommendations 1988. Biochemical Journal 258, 1–2. Ockenden, I., Dorsch, J.A., Reid, M.M., Lin, L., Grant, L.K., Raboy, V. and Lott, J.N.A. (2004) Characterization of the storage of phosphorus, inositol phosphate and cations in grain tissues of four barley (Hordeum vulgare L.) low phytic acid genotypes. Plant Science 167, 1131–1142. Parthasarathy, R. and Eisenberg, F. Jr (1991) Biochemistry, stereochemistry, and nomenclature of the inositol phosphates. In: Reitz, A.B. (ed.) Inositol Phosphates and Derivatives. American Chemical Society, Washington, DC, pp. 1–19. Sarma, N.S. (2004) Etymology as an aid to understanding chemistry concepts. Journal of Chemical Education 81, 1437–1439. Shears, S.B. (2004) How versatile are inositol phosphate kinases? Biochemical Journal 377, 265–280. Shears, S.B. (2005) Telomere maintenance by intracellular signals: new kid on the block? Proceedings of the National Academy of Sciences of the United States of America 102, 1811–1812. Tavares, P., Martinez-Salgado, C., Ribeiro, C.A., Elono, N., Lopez-Novoa, J.M. and Teixeira, F. (2002) Cyclosporin effect on rat aorta α1-adrenoceptors and their transduction mechanisms. Journal of Cardiovascular Pharmacology 40, 181–188. Torres, J., Domínguez, S., Cerdá, F.M., Obal, G., Mederos, A., Irvine, R.F., Dìaz, A. and Kremer, C. (2005) Solution behaviour of myo-inositol hexakisphosphate in the presence of multivalent cations. Prediction of a neutral pentamagnesium species under cytosolic/nuclear conditions. Journal of Inorganic Biochemistry 99, 828–840. Woodcock, E.A., Mitchell, C. J. and Biden, T. J. (2003) Phospholipase Cδ1 does not mediate Ca2+ responses in neonatal rat cardiomyocytes. FEBS Letters 546, 325–328. Yoshida, K.T., Wada, T., Koyama, H., Mizobuchi-Fukuoka, R. and Naito, S. (1999) Temporal and spatial patterns of accumulation of the transcript of myo-inositol-1-phosphate synthase and phytin-containing particles during seed development in rice. Plant Physiology 119, 65–72.

2

Identification of Inositol Phosphates by Nuclear Magnetic Resonance Spectroscopy: Unravelling Structural Diversity Pushpalatha P.N. Murthy

Department of Chemistry, Michigan Technological University, 1400 Townsend Drive, Houghton, MI 49931, USA

At first glance, inositols are deceptively simple molecules. On closer examination, a host of stereochemical, regiochemical, prochiral and conformational issues reveal themselves (Posternak, 1965; Parthasarathy and Eisenberg, 1986, 1990). In fact, the International Union of Pure and Applied Chemistry (IUPAC) needed three attempts and 26 years to agree on a system of nomenclature that adequately represents the stereochemical issues involved (IUPAC Commission on the Nomenclature of Organic Chemistry and IUPAC–IUB Commission on Biochemical Nomenclature (CBN), 1976; IUB Nomenclature Committee, 1989). The complexity is due to the presence of numerous stereochemical elements in the molecule, including nine stereoisomers of the parent inositol moiety (scyllo-, neo-, muco-, etc.), multiple phosphorylated derivatives (63 different compounds are possible in the case of myo-inositol) and the presence of conformational isomers (Posternak, 1965; Parthasarathy and Eisenberg, 1986, 1990; Murthy, 2006; see Shears and Turner, Chapter 1, this volume). A complete structural analysis of inositol phosphates therefore requires that all of these elements be determined. Structural analysis of inositol phosphates requires the extraction of polar, highly charged molecules with minimal structural perturbation, chromatographic purification and, finally, establishment of their molecular architecture. As inosi-

tol phosphates can carry numerous negative charges and often exist in chelated forms, their extraction and purification poses many challenges. Structures of inositol phosphates can be established by chemical degradation or nuclear magnetic resonance (NMR) spectroscopy (reviewed in Irvine, 1986). The procedure for chemical degradation involves subjecting a purified and radiolabelled inositol phosphate to a series of chemical and enzymatic reactions, followed by identification of the products by co-migration with standards (Fig. 2.1; Grado and Ballou, 1961). This is an elaborate and time-consuming method, which normally takes months to complete. In addition, it is an indirect method of structural determination and the conclusions are therefore ambiguous. In contrast, NMR spectroscopy is a versatile method that can provide complete structural information in a few hours (Derome, 1987; Friebolin, 1993; Claridge, 1999). Some of the advantages of NMR spectroscopy over conventional methods are: ●

●

Inositol phosphates contain three NMRactive nuclei (1H, 31P and 13C), so a detailed picture of the molecule can be gleaned by combining information from all nuclei. Structural conclusions are direct and unambiguous and are obtained without the need for co-migration with standards.

©CAB International 2007. Inositol Phosphates: Linking Agriculture and the Environment (eds B.L. Turner, A.E. Richardson and E.J. Mullaney)

7

8

P.P.N. Murthy

CH2OH P

P

P 5 1

HO

P OH OH

5

Periodate 4

OH

1. Reduction 2. Dephosphorylation

OH

1

4 HO

HH 2

2 P

OH

P O

OH

O CH2OH

Ins(1,4,5)P3

D-Iditol

Fig. 2.1. Chemical degradation of myo-inositol 1,4,5-trisphosphate (Ins(1,4,5)P3) to D-iditol.

●

●

●

Information can be obtained about the structure of the parent inositol moiety, the number and positions of phosphorylation on the ring, and the conformation of the cyclohexane ring. Dynamic processes such as conformational inversion and chemical reactions of inositol phosphates can be investigated. Analysis is non-destructive and involves minimal sample manipulation.

Disadvantages with NMR spectroscopy include: ●

●

●

Low sensitivity: about 0.1 µmol of the inositol phosphate is required, which corresponds to ~25 µg or 1 ml of ~100 µM sample. Inability to distinguish between enantiomers: additional experiments with shift reagents can provide enantiomeric information, but there are currently no established methods for inositol phosphates. The equipment is expensive and the technique is not user-friendly.

NMR spectroscopy has nevertheless been used to obtain a variety of information about inositol phosphates (Table 2.1). Many of these applications are reviewed here. This chapter is not meant to be complete or exhaustive, but includes discussion of the procedures most likely to be useful for the investigation of environmental samples.

Conformational Inversion Conformational isomers are structural isomers that are interconverted by rotation around single

bonds (Carey and Sundberg, 2000). The example of myo-inositol hexakisphosphate is shown in Fig. 2.2. The properties of conformational isomers, including size, shape, energy, chemical reactivity, ability to chelate with metal ions and binding interactions with proteins, can be markedly different. Conformational flexibility of biomolecules has a major impact on binding interactions with enzymes and receptors, and therefore on biological activity. The energy required for rotation around single bonds is low, with the activation energy for the chair– chair transition of cyclohexane being about 45 kJ/mol (10.8 kcal/mol) (Carey and Sundberg, 2000). This means that interconversion between conformational isomers occurs readily at room temperature and multiple low-energy conformers exist. Information on possible low-energy conformers at room temperature is therefore necessary to understand binding interactions with proteins and metal ions. NMR spectroscopy has been applied extensively to investigate the conformational flexibility and chair– chair interconversions of a number of inositol phosphates ( Johnson and Tate, 1969; Costello et al., 1976; Isbrandt and Oertei, 1980; Emsley and Niazi, 1981; Lasztity and Lasztity, 1990). In fact, the conformation adopted by myoinositol hexakisphosphate has been the subject of much debate and was one of the first applications of 31P NMR spectroscopy for structural investigation of inositol phosphates. Johnson and Tate (1969) employed 31P NMR spectroscopy to confirm the structure of myo-inositol hexakisphosphate and suggested that the conformation was the sterically favourable 1-axial/5-equatorial form (Fig. 2.2). However, X-ray crystal data of the

Identification by NMR Spectroscopy

9

Table 2.1. Applications of nuclear magnetic resonance (NMR) spectroscopy for the investigation of inositol phosphates. Application

Representative references

Conformational analysis, including activation energy of ring inversion of inositol tetrakis-, pentakis- and hexakisphosphates

Johnson and Tate (1969), Costello et al. (1976), Isbrandt and Oertei (1980), Emsley and Niazi (1981), Lasztity and Lasztity (1990), Brigando et al. (1995), Barrientos and Murthy (1996), Bauman et al. (1999), Paton et al. (1999), Blum-Held et al. (2001), Volkmann et al. (2002) Lemieux et al. (1957, 1958), Brownstein (1959), Dorman et al. (1969), Angyal et al. (1974), Angyal and Odier (1982), Cerdan et al. (1986), Lindon et al. (1986), Mayr and Dietrich (1987), Szwergold et al. (1987), Hansen et al. (1989), Barrientos et al. (1994), Johnson et al. (1995), Barrientos and Murthy (1996) Barrientos et al. (1994), Johnson et al. (1995), Raboy et al. (2000), Dorsch et al. (2003)

Structural determination of pure inositol phosphates

Structural determination of multiple inositol phosphates in a mixture without separation Structural determination of impure samples without separation Acid dissociation constants (pKa) and protonation sequences of inositol phosphates at the microscopic level Intramolecular hydrogen bonding in inositol phosphates Support for theoretical calculations Solid-state NMR spectroscopy

Volkmann (2002) Schmitt et al. (1993), Brigando et al. (1995), Schlewer et al. (1999), Blum-Held et al. (2001), Borkovec and Spiess (2004)

Felemez and Spiess (2003) Bauman et al. (1999), Volkmann et al. (2002), Yang et al. (2005) Gardiennet et al. (2005)

dodecasodium salt clearly indicated that in solid state it was in the sterically unfavourable 5-axial/ 1-equatorial form (Blank et al., 1971). A number of subsequent NMR studies suggested that notwithstanding bulky phosphate groups, conformational inversion to the sterically hindered form does indeed occur in myo-inositol hexakisphosphate and other inositol phosphates (Isbrandt and Oertei, 1980; Emsley and Niazi, 1981; Brigando et al., 1995; Barrientos and

O− −O

O− P 4 O O O −O O P O O O−

O−

P 3

O

O P O−

O−

O

1

O

−O

P

−O

−O

O P

4

2

O

O O−

P

P O

O

O− −O

O

O−

−O

Murthy, 1996; Bauman et al., 1999; Paton et al., 1999; Blum-Held et al., 2001; Volkmann et al., 2002). Numerous theoretical studies also support these conclusions (Bauman et al., 1999; Volkmann et al., 2002; Yang et al., 2005). However, the pH dependence of the conformational inversion process and the structural and environmental factors that contribute to stabilizing the sterically hindered form are not completely understood. The fact that the coupling constants in 1H-NMR

O− P

O

O

O−

O O

P −O

1-axial/5-equatorial

−O

P

1 O

O 3

−O

O P 2

−O

O−

O− 5-axial/1-equatorial

Fig. 2.2. Conformational interconversion of myo-inositol hexakisphosphate.

O

10

P.P.N. Murthy

provide information about the dihedral angles between vicinal protons (coupling constants with vicinal protons) has been particularly useful in the investigation of chair– chair interconversions (Barrientos and Murthy, 1996). In addition, dynamic NMR spectroscopy has been used to provide information on the activation energy of ring flipping, while two-dimensional random delay exchange spectroscopy (EXSY) has been applied to observe the interconversion of molecules between different conformations (Fig. 2.3; Bauman et al., 1999; Volkmann et al., 2002). These investigations clearly indicated that myoinositol hexakisphosphate, scyllo-inositol hexakisphosphate, all isomers of myo-inositol pentakisphosphate and one isomer of myo-inositol tetrakisphosphate undergo ring– ring interconversion from the 1-axial/5-equatorial form to the 5-axial/1-equatorial form at room temperature. Extensive application of 1H, 31P and twodimensional NMR techniques, supported by theoretical studies, has indicated that the energy of conformations is influenced by four factors:

4/6

1/3 5

2 F2 (ppm)

4/6 1/3 5

2

3.4 3.5

(i) number, substitution pattern and stereochemistry of phosphate groups on the inositol backbone; (ii) stereochemistry of the parent inositol ring; (iii) physical state (solid or aqueous solution) of the compound; and (iv) properties of the solvent, such as pH and counter-ions. From these studies, the following generalizations can be made (Barrientos and Murthy, 1996; Bauman et al., 1999; Paton et al., 1999; Blum-Held et al., 2001; Volkmann et al., 2002): ●

●

●

●

The pH-dependent conformational preferences of inositol phosphates are unique to the particular isomer and do not parallel the behaviour of myo-inositol hexakisphosphate. The presence of four or more equatorial phosphates on the inositol ring induces a change in the conformation from the sterically unhindered 1-axial/5-equatorial structure to the sterically hindered 5-axial/1-equatorial conformation at high pH (Fig. 2.4). myo-Inositol hexakis- and pentakisphosphates exist in the 5-axial/1-equatorial conformation at pH > 10 (Fig. 2.4). myo-Inositol 1,4,5,6-tetrakisphosphate, which contains four contiguous equatorial phosphate groups, undergoes conformational inversion to the sterically hindered 5-axial/1 equatorial form. However, myo-inositol 1,2, 3,4-tetrakisphosphate and myo-inositol 1,2,5,6tetrakisphosphate do not (Fig. 2.5).

3.6 3.7 3.8

Structural Determination of Purified Inositol Phosphates

3.9 4.0 4.1 4.2 4.3 4.4 4.5 4.6 4.7 4.6 4.5 4.4 4.3 4.2 4.1 4.0 3.9 3.8 3.7 3.6 3.5 3.4 F1 (ppm)

Fig. 2.3. Random delay exchange spectroscopy (EXSY) of myo-inositol hexakisphosphate at pH 9.2 with attached 1H spectra. Protons of the 1-axial/5-equatorial conformer are indicated above the resonances and protons of the 5-axial/ 1-equatorial form below. The sweep width in both the F1 and F2 dimensions was 1355.7 Hz.

The application of NMR spectroscopy to the structural determination of inositols dates back to the early days of NMR spectroscopy in the 1950s (Lemieux et al., 1957, 1958; Brownstein, 1959). Those early investigations quickly revealed that NMR spectroscopy could distinguish between axial and equatorial groups, because axial protons occur at higher field positions compared to equatorial protons. They also revealed that the chemical shifts and coupling constants of protons are influenced by the molecular configuration, so NMR techniques held great promise for analysing the configuration and conformation of inositol phosphates (Lemieux et al., 1957, 1958;

Identification by NMR Spectroscopy

P

P P

P

P

P

2 P

P

P

1

1

P

11

2

P P

P

P

P

P

P

P

P

P

P P

P

P

P

P

P

P

2 P 1

P

P 1 P

P

2

P

P

OH

P

2 P

P HO

1

OH P

OH 2

P

P

HO

P

P

P

P

P

P

P OH 2 P 1

P

1

1

P

P 2

P

Fig. 2.4. Conformational inversion of myo-inositol pentakis- and hexakisphosphates.

P

P

P

P P

2

P

HO

P

P

OH

1

1

OH OH

2

Ins(1,2,3,4)P4

P

OH P

HO

OH

P P

2 P

P P

1

OH

1

2

P

Ins(1,2,5,6)P4 P

OHOH

P OH

2

P P

P P

1

1

P

OH 2

P

Ins(1,4,5,6)P4 Fig. 2.5. Conformational inversion of myo-inositol tetrakisphosphates.

Brownstein, 1959). Investigation of the 1H and 13 C NMR spectra of all nine stereoisomers of inositol clearly demonstrated that those with three axial groups (cis-, allo- and muco-) undergo chair–chair interconversion at room temperature (Dorman et al., 1969; Angyal et al., 1974; Angyal and Odier, 1982). The interconversion of cisinositol is slower than the allo- and muco-forms, which interconvert rapidly at room temperature. It was hypothesized that this could be because each hydroxyl group in cis-inositol has to pass between two hydroxyl groups during the process (Angyal and Odier, 1982). In the 1980s, NMR spectroscopy (1H, 13C and 31P) was employed to determine the structure of naturally-occurring inositol phosphates isolated and purified from cells (Cerdan et al., 1986; Lindon et al., 1986; Mayr and Dietrich, 1987; Szwergold et al., 1987; Hansen et al., 1989). These studies established the usefulness of NMR spectroscopy for the structural elucidation of myo-inositol phosphates, including myo-inositol 1,4,5-trisphosphate, inositol tetrakisphosphates and myo-inositol hexakisphosphate, and established NMR parameters such as chemical shifts, coupling constants and multiplicity patterns for structural analysis of 1 H, 13C and 31P NMR spectra of inositol phosphates. The principal NMR parameters that pro-

12

P.P.N. Murthy

vide structural information include the following (Barrientos et al., 1994; Johnson et al., 1995; Barrientos and Murthy, 1996): ●

●

●

●

The number of chemically distinct sets of resonances. The presence of a plane of symmetry would result in four distinct sets of resonances (or fewer if resonances overlap as in myo-inositol hexakisphosphate) and six if there were no plane of symmetry. Chemical shifts of nuclei. These are influenced by the electronic environment, such as the axial or equatorial orientation of the proton, the nature and number of geminal and vicinal substituents, and the ionization state. The lack of a phosphate group results in the upfield shift of the α-proton by about 0.5–1.0 ppm. Multiplicity and coupling constants of resonances. These provide information about the number and orientation of vicinal protons, dihedral angles and the presence of geminal phosphates. The splitting pattern of protons on the inositol ring is due to coupling with two vicinal protons, one on either side (Jax–eq about 2–3 Hz, Jeq–eq about 2–3 Hz, Jax–ax about 8–10 Hz) and with the phosphorus ( JH–P about 8–10 Hz). Therefore, the presence of a phosphate group significantly affects the splitting pattern of the inositol ring proton, due to the additional 8–10 Hz 1 H–31P coupling. Depending on the structure, long-range coupling (W coupling) can also be detected (Cerdan et al., 1986; Barrientos et al., 1994). Two-dimensional NMR experiments. These provide a wealth of information about connectivity and dynamic processes (Derome, 1987; Friebolin, 1993; Claridge, 1999). The

application of these general principles for the structural determination of inositol phosphates is elaborated in the following section.

Structural Determination of Individual Inositol Phosphates in a Mixture without Purification: Application of Two-dimensional Total Correlation Spectroscopy Cells generally contain multiple inositol phosphates. The metabolism of inositol phosphates is tightly interconnected by the action of fast-acting phosphatases and kinases. Therefore, to get a complete picture of the metabolism of inositol phosphates, changes in the concentration of multiple inositol phosphates must be monitored simultaneously. Purification of individual inositol phosphates from a mixture is neither easy nor always possible. Analysis of NMR spectra of a mixture containing multiple components is difficult due to the inability to assign resonances to individual molecules, especially in regions where multiple resonances overlap. An NMR technique that allows the structural assignment of individual components in a mixture without prior separation would greatly simplify the problem of structural determination. Two-dimensional total correlation spectroscopy (TOCSY) experiments can be used to determine networks of mutually coupled protons (Derome, 1987; Griesinger et al., 1988; Claridge, 1999, pp. 201–211). With the addition of a spin lock period the magnetization of H1 is transferred to H2, H3 and H4; i.e. magnetization is relayed down a chain of contiguous spin-coupled protons past the vicinal protons (Fig. 2.6). In a

TOCSY

H

H1

H2

H3

H4

O

C

C

C

C

C

H5 X

C

OH OH

H P

HO

H OH HO

H

H H Fig. 2.6. Coupling pathway for myo-inositol 5-monophosphate mapped by a TOCSY sequence.

Identification by NMR Spectroscopy

13

Determination of myo-inositol phosphates in a mixture

two-dimensional spectrum, H2 will show cross peaks to H1, H3 and H4. All protons on each inositol phosphate are part of a connected spin system. Therefore, all protons should show connectivity either due to direct coupling or longrange magnetization relay. Thus, the TOCSY technique provides a way of identifying all the resonances belonging to individual inositol phosphates ( Johnson et al., 1995). In addition, individual 1H spectra of each component can be extracted from two-dimensional TOCSY data (Barrientos et al., 1994; Johnson et al., 1995; Raboy et al., 2000; Dorsch et al., 2003). Two examples of the use of TOCSY experiments to determine the structure of individual inositol phosphates in a mixture of inositol phosphates are described below.

The sequential hydrolysis of myo-inositol hexakisphosphate by phytase produces multiple inositol phosphates. Figure 2.7 shows the TOCSY spectrum of a mixture of inositol phosphates obtained by alkaline phytase catalysed hydrolysis of myoinositol hexakisphosphate. From the one-dimensional proton spectrum (top) it is not possible to confidently deduce either the number or the structures of inositol phosphates in the mixture. However, the two-dimensional TOCSY spectrum suggests the presence of three spin systems (i.e. three inositol phosphates) indicated by horizontal lines [H], [I] and [ J], with several overlapping resonances. To illustrate the interpretation of a

P P 2

P P

P

[J]

P

1

F1 (ppm)

4

2

3.6

3 1

6

5 [H] [I]

1,3

4,6

2

5

3.8

P P 2

P

[I]

4.0

P

HO P

1 4.2

[H]

[J]

4.4

P P

4,6

2

1,3,5

2

P

P

HO HO

4.6

1 4.8

4.8

4.6

4.4

4.2

4.0

3.8

3.6

3.4

F2 (ppm)

Fig. 2.7. Two-dimensional TOCSY spectrum of a mixture of inositol phosphates obtained after 2 h of alkaline phytase–catalysed hydrolysis of myo-inositol hexakisphosphate (Johnson et al., 1995). Proton spectra are attached. The sweep width in both F1 and F2 dimensions was 1084 Hz. Horizontal lines have been drawn to indicate resonances that arise from molecules [H], [I] and [J]. Structures and proton assignments of [H], [I] and [J] are shown.

14

P.P.N. Murthy

TOCSY spectrum, Fig. 2.7 is discussed in detail below. One compound gives rise to six peaks as indicated by the arrow [H], a second gives rise to four peaks as indicated by the arrow [I] and a third gives rise to three peaks as indicated by the arrow [ J]. The presence of three sets of resonances in [ J] suggests a plane of symmetry in the inositol phosphate as well as overlapping resonances. The relative downfield chemical shifts of all the 1H resonances suggest that all the carbons are phosphorylated, so the compound must be myo-inositol hexakisphosphate. The chemical shifts, multiplicity, coupling constants and overlap of signals from protons in the H-1, H-3 and H-5 positions of the inositol ring provide additional evidence of this assignment. The spectrum of this compound is well documented and has been discussed in detail (e.g. Barrientos and Murthy, 1996). The presence of four sets of resonances in [I] indicates a symmetrical molecule. The most noticeable change in [I] compared to [ J] is the upfield shift of one resonance by d ~0.8 ppm to d 3.6 ppm, which indicates that dephosphorylation has occurred on one of the carbons in the plane of symmetry (i.e. either C-2 or C-5). The presence of a triplet at d 3.6 ppm rather than a quartet confirms the loss of 1H–31P coupling. The equatorial proton at H-2 has a characteristic resonance at ~d 4.8 ppm (a triplet with J ~ 2–3 Hz), so the upfield shifted of one by ~0.5 ppm to d 3.8 ppm resonance. The coupling constants of ~8 Hz also provide additional evidence that the resonance at d 3.6 ppm is due to H-5, because H-2 would give rise to a triplet with J of ~2 Hz. Therefore, [I] must be myo-inositol 1,2,3,4,6-pentakisphosphate. The presence of six sets of resonances in [H] indicates a lack of symmetry in the molecule. The most noticeable change in [H] compared to [I], is the upfield shift of one resonance by about d 0.5–3.8 ppm. The triplet splitting pattern ( J ~ 8 Hz) indicates the loss of the 1H–31P coupling and suggests that the resonance must be a proton at H-6 (or H-4) and not H-1 (or H-3), which would give rise to a doublet ( J = 8–9 Hz and 2–3 Hz). Therefore, [H] must be a tetrakisphosphate, either myo-inositol 1,2,3,4-tetrakisphosphate or the 1,2, 3,6 enantiomer. The connectivity indicated by [H], [I] and [ J] is repeated several times in the

spectrum, both horizontally and vertically, because all the protons of the inositol ring are connected and therefore show the same connectivity pattern. If the composite spectrum is complicated with several overlapping resonances, the structures of the inositol phosphates can be determined by extracting the sub-spectra from the twodimensional TOCSY data (Johnson et al., 1995).

Inositol phosphates in plant seeds The TOCSY experiment was employed to investigate the inositol phosphate phenotype of mutant barley and maize seeds (Raboy et al., 2000; Dorsch et al., 2003). Seeds contain a complex mixture of highly phosphorylated inositol phosphates, so the separation and structural determination of inositol phosphates in such samples pose a formidable challenge. The 1H-NMR spectra of inositol phosphates in wild-type barley seeds are shown in Fig. 2.8. The one-dimensional 1H-NMR spectrum on top of Fig. 2.8 is complex and contains many overlapping resonances. Therefore, it was not possible to confidently deduce either the number or the structures of inositol phosphates in the mixture. The two-dimensional TOCSY spectrum, however, revealed four sets of mutually coupled spin systems (i.e. four inositol phosphates) labelled [11], [12], [14] and [17]. Consideration of the chemical shifts, coupling constants and multiplicity patterns of each spin system, as described above, helped establish the structures as myo-inositol hexakisphosphate, myoinositol 1,2,3,4,6-pentakisphosphate, myo-inositol 1,2,3,5,6-pentakisphosphate (or its enantiomer myo-inositol 1,2,3,4,5-pentakisphosphate) and myoinositol 1,2,4,5,6-pentakisphosphate (or its enantiomer myo-inositol 2,3,4,5,6-pentakisphosphate) (Dorsch et al., 2003). The assignment of resonances is indicated in the figure. Thus, the structures of inositol phosphates in a mixture containing four highly phosphorylated derivatives of myo-inositol were readily ascertained without separation. In summary, a two-dimensional TOCSY experiment obviates the need for chromatographic separation and provides sufficient information to unambiguously assign the structures of closely related compounds in about 3 or 4 h rather than months.

Identification by NMR Spectroscopy

H2

15

H4,H6 H1,H3/H5 H4 H1,H3 H4

H5

H6

H4,H6

[17] H6

[11]

H3,H5

H1

[14/15]

H1,H3

H5

[12/13]

P P 2

P P

P

[17]

1

P

5.0

4.8

4.6

4.4

4.2

4.0

3.8

3.6

ppm

P P

[11] 2

P

[14/15] [12/13]

[17]

[12] P

HO 1

P

F2 (ppm) 3.6

5

P P P

2

HO

[11] P

P

1

3.8

1

6 4.0 5

4.2 4.4

OH

1,3/5

1,3

4,6

4

3,5 1,3 6

4,6

4

4.6 P

P

P

P P

[14]

2

4.8 2 5.0 5.0

4.8

4.6

4.4

4.2

4.0

3.8

3.6

3.4

F1 (ppm)

Fig. 2.8. One-dimensional 1H spectrum (top) and two-dimensional TOCSY spectrum (bottom) of a mixture of inositol phosphates extracted from wild-type barley seeds (Dorsch et al., 2003). Proton spectra are attached. Vertical lines have been drawn to indicate resonances that arise from molecules [11], [12], [14] and [17], and the structures and proton assignments of the molecules are shown.

Structural Determination of Impure Samples with Complex Proton Spectra When analysing samples that are impure and/or display crowded proton resonances, the TOCSY experiment may provide spectra that are still too crowded for unambiguous interpretation. In such situations, the presence of the NMR-active heteroatom 31P (natural abundance 100%) in inositol phosphates can be used as a means to pull out

the 1H spectrum of inositol phosphates. The proton–proton connectivity information can be sorted by the 31P chemical shift attached to the network. Thus, the addition of a TOCSY spin lock mixing period after the heteronuclear multiple quantum correlation (HMQC) sequence allows magnetization transfer on to neighbouring protons (Fig. 2.9). The result is a 31P-selected two-dimensional TOCSY spectrum; in other words, only the proton resonances attached to phosphorus-containing molecules are pulled out

16

P.P.N. Murthy

TOCSY

H

H1

H2

H3

H4

O

H5

C

C

C

C

C

C

OH OH

H P

HMQC

H

HO OH HO

P

H

H

H

Fig. 2.9. Coupling pathway for myo-inositol 5-monophosphate mapped by a heteronuclear multiple quantum correlation–total correlation spectroscopy (HMQC–TOCSY) sequence.

of the complex 1H-NMR spectrum (Friebolin, 1993; Braun et al., 1998; Claridge, 1999, pp. 241–243). As an example, the HMQC–TOCSY spectrum of a mixture of myo-inositol hexakisphosphate and unphosphorylated myo-inositol is shown in Fig. 2.10. The resonances downfield of d 4.2 ppm (F2-axis) due to myo-inositol hexakisphosphate are

H

H

P

P H P

P P P

H

HO HO HO

2 OH

P(4,6)

1

P(1,3) P(2)

P(5)

F2 (ppm) 3.2

C

H(1,3)

OH

H

H(5)

H

3.4 3.6

4.0 4.2 4.4 4.6

H(2)

InsP6

A

3.8

H(1,3,5) H(4,6)

H(2)

H(4,6)

Ins

HO

coupled to phosphorus nuclei (on the F1-axis). In contrast, the resonances upfield of d 4.0 ppm due to myo-inositol are not coupled to phosphorus nuclei. Line A indicates that the phosphate at the P-2 position of myo-inositol hexakisphosphate is coupled to all the protons on the inositol ring, as these protons are mutually coupled. This is also true for P-1,3, P-4,6 and P-5. Line B shows that

4.8

B

5.0 5.2 5.4 9.0

8.5

8.0

7.5 F1 (ppm)

7.0

6.5

6.0

Fig. 2.10. Two-dimensional HMQC–TOCSY spectrum of a mixture of myo-inositol hexakisphosphate and myo-inositol with attached one-dimensional 31P (top) and 1H (left) spectra (Volkmann, 2002). Vertical and horizontal lines have been drawn to indicate coupled resonances, and the resonances due to each compound are indicated.

Identification by NMR Spectroscopy

17

constants of the 1H resonances and the information on 31P coupling allow the unambiguous assignment of the molecular structures. This technique holds great promise for the analysis of complicated mixtures of inositol phosphates, as well as in vivo NMR spectroscopy. These applications are currently under investigation.

the proton at the H-2 position is coupled to all four 31P resonances, indicating that H-2 is connected to a molecule with all four phosphates. This is also the case with H-4,6 and H-1,3,5. The inositol molecule does not contain any 31P and therefore does not appear in the contour map; thus the complete proton spectrum of molecules to which the 31P is attached is provided (Volkmann, 2002). Figure 2.11 shows the spectrum of a mixture of myo-inositol hexakisphosphate, myo-inositol and glucose 6-phosphate. The 1H-NMR spectrum on the left is complicated; it contains many overlapping resonances and provides insufficient information to make structural assignments. The HMQC–TOCSY spectrum clearly indicates the presence of one molecule with one phosphate (line D, glucose 6-phosphate), a second with four phosphates (line E, myo-inositol hexakisphosphate) and one not coupled to any 31P resonances (myoinositol). The chemical shifts and the coupling

Protonation Sequence at Microscopic Level, Acid Dissociation Constants and Hydrogen Bonding Potentiometric studies provide acid dissociation constants (pKa) at a macroscopic level, as well as overall protonation or dissociation constants that describe the molecule as a whole. In inositol phosphates, all phosphates are not equivalent. Therefore, the microscopic pKa values differ and potentiometric measurements do not provide

P(5)

P(4,6) P(1,3) P(2)

Glu-6-P D

F2 (ppm) 3.0 3.2 3.4

Ins

3.6 3.8 4.0 4.2 4.4

InsP6

4.6 4.8

E

5.0 5.2 5.4 10.5

10.0

9.5

9.0

8.5

8.0 7.5 F1 (ppm)

7.0

6.5

6.0

5.5

5.0

Fig. 2.11. Two-dimensional HMQC–TOCSY spectrum of a mixture of myo-inositol hexakisphosphate, glucose 6-phosphate and myo-inositol with attached one-dimensional 31P (top) and 1H (left) spectra (Volkmann, 2002). Vertical and horizontal lines have been drawn to indicate coupled resonances, and the resonances due to each compound are indicated. 1H resonances of glucose 6-phosphate are not indicated because they partially overlap with the myo-inositol resonances as indicated by line D. The 31P resonances of myo-inositol hexakisphosphate and glucose 6-phosphate are indicated on the F1 axis. Lines D and E indicate coupled resonances.

18

P.P.N. Murthy

information on the ionization state of individual phosphates. As the chemical shift of 31P is influenced mainly by the electronic effects that accompany protonation and deprotonation, the protonation sequence at a microscopic level (i.e. the sequence of protonation or deprotonation of the various phosphate groups) can be determined by monitoring the change in the chemical shift of 31 P as a function of pH. The protonation sequence of myo-inositol hexakisphosphate, myoinositol 1,4,5-trisphosphate and a number of other inositol phosphates has been investigated by 31P NMR spectroscopy. Numerous studies have assessed the pKa values of individual phosphates on inositol phosphates (Brigando et al., 1995; Schlewer et al., 1999; BlumHeld et al., 2001; Borkovec and Spiess, 2004). For myo-inositol hexakisphosphate, Brigando et al. (1995) combined data from potentiometric studies with NMR studies to suggest that the approximate pKa values of the 12 protons are as follows: the first proton on the phosphates at the P-2, P-5 and P-1,3 positions are the most acidic, with pKa values less than 2; P-4,6 are less acidic, with pKa values of 2.6. The pKa values of the second protons of P-5 and P-2 are 6 and 7, respectively, while those of P-1,3 and P-4,6 are ~9, 10, 11 and 12, respectively. A similar study was undertaken to determine the microionization constants of myo-inositol tris- and tetrakisphosphates (Schmitt et al., 1993; Schlewer et al., 1999; BlumHeld et al., 2001; Borkovec and Spiess, 2004), and the sequence of deprotonation of these compounds has been elucidated in great detail. These investigations allow us to pinpoint the exact location of negative charges at a given pH. Recently, 1H-NMR methods were employed to investigate hydrogen-bonding interactions in inositol phosphates. Felemez and Spiess (2003) monitored the change in chemical shift of hydroxyl protons as a function of pH and suggested the formation of an intramolecular hydrogen bond between the 1-hydroxyl and 2-phosphate in myoinositol 2-monophosphate (Fig. 2.12).

Analysis of Inositol Phosphates in Environmental Samples by Nuclear Magnetic Resonance Spectroscopy Although the presence of inositol phosphates in terrestrial and aquatic ecosystems has been known

O−

O P

OH

O

O

O

P H

O

O− H

O

O H

[I]

[II]

Fig. 2.12. Intramolecular hydrogen-bonding interactions in myo-inositol 1,2,6-trisphosphate. [I] and [II] represent alternative intramolecular hydrogen-bonding structures between monoprotonated phosphates and vicinal hydroxyl groups.

for a long time, little is known about the composition, cycling, mobility or bioavailability of inositol phosphates in the environment (reviewed in Turner et al., 2002). The environmental concerns raised by agricultural phosphate contamination, particularly in areas of high livestock density (see Leytem and Maguire, Chapter 10, this volume), have highlighted the need to accurately monitor inositol phosphates in soils and aquatic sediments. However, the analytical difficulties associated with these studies pose a major challenge. The extraction and purification of inositol phosphates in soil is complicated by their complexation with polyvalent metal ions and association with complex organic matter such as humic acids (see Celi and Barberis, Chapter 13, this volume). In addition, soils contain uncommon phosphorylated stereoisomers of inositol (scyllo-, neo- and D-chiro-) in abundance (see Turner, Chapter 12, this volume). Methods for the efficient extraction, purification and structural assignment of organic phosphates in soils and other environmental samples have been explored and significant advances have been made recently (Turner et al., 2002; Turner and Richardson, 2004; see Cooper et al., Chapter 3, this volume). The use of one-dimensional 31P NMR spectroscopy to analyse alkaline soil extracts indicated that scyllo-inositol hexakisphosphate, a compound not reported in plants and animals to date, is a major component of the soil organic phosphorus (Turner and Richardson, 2004; Turner, Chapter 12, this volume). The inclusion of a hypobromite oxidation step, which destroys all organic phosphates except the inositol

Identification by NMR Spectroscopy

phosphates, can significantly help structural assignment. Structural identification using one-dimensional 31P NMR spectroscopy has several limitations, including low sensitivity of 31P compared with 1 H (6%), narrow spread and poor resolution of phosphate resonances in the phosphate monoester region, and the singlet multiplicity of phosphate resonances that does not provide structural information about the molecular environment of phosphates (i.e. other nuclei to which the phosphates are coupled). The latter is of particular concern, because the presence of rare inositol stereoisomers in soil, such as scyllo-, neo- and D-chiro-inositol, requires structural information about the inositol ring connected to phosphates for complete structural identification. Some of these limitations may be overcome by using two-dimensional techniques such as HMQC–TOCSY discussed above. This method may also eliminate the need for hypobromite oxidation and the potential structural changes associated with it. For studies involving the concentration and movement of inositol phosphates in the environment, solid-state NMR would be the preferred method of investigation, but the technique presents many problems for application to soils. These include reduced sensitivity and line broadening, the presence of paramagnetic ions, narrow spread of 31P resonances so structural information is hard to obtain and the general difficulty in extracting information from solid-state NMR (Condron et al., 1997). An interesting example of the use of solid-state NMR spectroscopy for inositol phosphates was recently described by Gardiennet et al. (2005), who used solid-state 31P NMR spectroscopy to detect the presence of the mono- and dianionic species of myo-inositol 2monophosphate.

Summary and Recommendations for Future Research In summary, 1H, 31P and 13C NMR experiments have revealed many structural details of inositol phosphates, but a number of challenges remain. The intrinsic low sensitivity of NMR spectroscopy and the low endogenous concentrations of inositol phosphates in cells make it difficult to monitor inositol phosphate metabolism in vivo. New techniques need to be developed for in vivo studies

19

and environmental samples. Development of methods based on 31P-selected two-dimensional methods such as HMQC–TOCSY appears promising for solution NMR. The need to minimize sample manipulation in soils means that the application of solid-state NMR methods to inositol phosphates in soil samples requires investigation.

Experimental Details This section provides additional details for the experiments discussed above. NMR spectra were recorded on a 400 MHz Varian Unity Inova-400 spectrometer. The samples were dissolved in deuterium oxide (0.8 ml) and the pH adjusted to 5.0 with the addition of 1 M NaOH or perdeuterated acetic acid, as necessary. Onedimensional 1H-NMR spectra were obtained at 399.943 MHz. 1H chemical shifts were referenced to the residual proton absorption of the solvent deuterium oxide (d 4.67 ppm). For onedimensional spectra, 16 scans with recycle delay of 6 s between acquisitions were collected. The acquisition conditions were as follows: spectral windows 5000 Hz; pulse width 90°. Typically, 16–32 scans were collected with recycle delays of 4–6 s between acquisitions. The residual water resonance was suppressed by a 2 s selective presaturation pulse. Two-dimensional EXSY was employed at 3°C and pH 10.7. The pulse sequence was a two-dimensional nuclear Overhauser effect spectroscopy (NOESY) with a missing time variation increment of 0.1 s (Claridge, 1999, pp. 326–328). A total of 128t1 increments were obtained, each consisting of four transients with a relaxation delay of 4 s between successive transients. A shifted Gaussian window was applied in both dimensions. The data matrix was expanded to a 1024 × 1024 real matrix. TOCSY data-sets were obtained with a 1H probe using the pulse sequence of Griesinger et al. (1988). Typically, 128t1 increments were collected, each consisting of 16–24 transients with a relaxation delay of 6 s between successive transients, using a TOCSY mixing time of 80 ms (as determined by one-dimensional TOCSY). The residual water signal was suppressed as in the one-dimensional experiment. A Laurentz–Gauss window was applied in both dimensions, and the data matrix was expanded to a 1024 × 512 real

20

P.P.N. Murthy

matrix. Digital resolutions in the F1 and F2 dimensions were ~4 and 2 Hz/point, respectively. HMQC–TOCSY parameters employed for the experiments were from Varian Instruments, ‘hmqctocsy’ (Varian, 1998). Typically, the parameters were as follows: the F1 dimension was 1448.9 Hz and F2 was 1600 Hz. A total of 270 t1 increments of 16 transients each were collected with a mixing time of 80 ms (as determined by one-dimensional TOCSY) and a relaxation delay

of 4 s. Residual water was suppressed as in the one-dimensional experiment with pre-saturation.

Acknowledgements The author thanks the National Science Foundation (Grant No. CHE-9512445) and Michigan Technological University for funds to purchase a 400 MHz NMR spectrometer.

References Angyal, S.G. and Odier, L. (1982) The 13C-NMR spectra of inositols and cyclohexanepentols: the validity of rules correlating chemical shifts with configuration. Carbohydrate Research 100, 43–54. Angyal, S.G., Greves, D. and Pickles, V.A. (1974) The stereochemistry of complex formation of polyols with borate and periodate anions, and with metal cations. Carbohydrate Research 35, 165–173. Barrientos, L.G. and Murthy, P.P.N. (1996) Conformational analysis of myo-inositol phosphates. Carbohydrate Research 296, 39–54. Barrientos, L.G., Scott, J.J. and Murthy, P.P.N. (1994) Specificity of phytic acid hydrolysis by alkaline phytase from lily pollen. Plant Physiology 106, 1489–1495. Bauman, A.T., Chateauneuf, G.M., Boyd, B.R., Brown, R.E. and Murthy, P.P.N. (1999) Conformational inversion process in phytic acid: NMR spectroscopic and molecular modeling studies. Tetrahedron Letters 40, 4489–4492. Blank, G.E., Pletcher, J. and Sax, M. (1971) The structure of myo-inositol hexaphosphate dodecasodium salt octatriacontahydrate: a single crystal X-ray analysis. Biochemical and Biophysical Research Communications 44, 319–325. Blum-Held, C., Bernard, P. and Spiess, B. (2001) myo-Inositol 1,4,5,6-tetrakisphosphate and myo-inositol 3,4,5,6tetrakisphosphate, two second messengers that may act as pH-dependent molecular switches. Journal of the American Chemical Society 123, 3399–3400. Borkovec, M. and Spiess, B. (2004) Microscopic ionization mechanism of inositol tetrakisphosphates. Physical Chemistry Chemical Physics 6, 1144–1151. Braun, S., Kolinowski, H.-O. and Berger, S. (1998) 150 and More Basic NMR Experiments: A Practical Course. WileyVCH, New York, pp. 505–508. Brigando, C., Mossoyan, J.C., Favier, F. and Benlian, D. (1995) Conformational preferences and protonation sequence of myo-inositol hexaphosphate in aqueous solution: potentiometric and multinuclear nuclear magnetic resonance studies. Journal of the Chemical Society, Dalton Transaction 4, 575–578. Brownstein, S. (1959) Shifts in nuclear magnetic resonance absorption due to steric effects. II. Polysubstituted cyclohexanes. Journal of the American Chemical Society 81, 1606–1608. Carey, F.A. and Sundberg, R. J. (2000) Conformational, steric, and stereoelectronic effects. In: Advanced Organic Chemistry, Part A, Structure and Mechanism, 4th edn. Kluwer Academic/Plenum Publishers, New York, pp. 123–185. Cerdan, S., Hansen, C.A., Johnson, R., Inubushi, T. and Williamson, J.R. (1986) Nuclear magnetic resonance spectroscopic analysis of myo-inositol phosphates including inositol 1,3,4,5-tetrakisphosphate. The Journal of Biological Chemistry 261, 14676–14680. Claridge, T.D.W. (1999) High Resolution NMR Techniques in Organic Chemistry. Pergamon Press, Oxford. Condron, L.M., Frossard, E., Newman, R.H., Tekely, P. and Morel, J.-L. (1997) Use of 31P NMR in the study of soils and the environment. In: Nanny, M.A., Minear, R.A. and Leenheer, J.A. (eds) Nuclear Magnetic Resonance Spectroscopy in Environmental Chemistry. Oxford University Press, New York, pp. 247–271. Costello, A.J.R., Glonek, T. and Myers, T.C. (1976) 31P nuclear magnetic resonance-pH titrations of myo-inositol hexaphosphate. Carbohydrate Research 44, 319–325. Derome, A.E. (1987) Modern NMR Techniques for Chemistry Research. Pergamon Press, Oxford. Dorman, D.E., Angyal, S.J. and Roberts, J.D. (1969) Nuclear magnetic resonance spectroscopy: 13C spectra of unsubstituted inositols. Proceedings of the National Academy of Sciences of the United States of America 63, 612–614.

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Dorsch, J.A., Cook, A., Young, K.A., Anderson, J.M., Bauman, A.T., Volkmann, C.J. and Murthy, P.P.N. (2003) Seed phosphorus and inositol phosphate phenotype of barley low phytic acid genotypes. Phytochemistry 62, 691–704. Emsley, J. and Niazi, S. (1981) The structure of myo-inositol hexaphosphate in solution: 31P NMR investigation. Phosphorus and Sulfur 10, 401–408. Felemez, M. and Spiess, B. (2003) 1H NMR titrations of hydroxy protons in aqueous solution as a method of investigation of intramolecular hydrogen-bonding in phosphorylated compounds: examples of myo-inositol 2phosphate and myo-inositol 1,2,6-tris(phosphates). Journal of the American Chemical Society 125, 7768–7769. Friebolin, H. (1993) Basic One- and Two-Dimensional NMR Spectroscopy. VCH, New York, pp. 287–305. Gardiennet, C., Henry, B., Kuad, P., Spiess, B. and Tekely, T. (2005) Straightforward detection of the secondary ionisation of the phosphate and pK determinations by high-resolution solid-state 31P NMR. Chemical Communications 180–182. Grado, C. and Ballou, C.E. (1961) myo-Inositol phosphates obtained by alkaline hydrolysis of beef brain phosphoinositide. Journal of Biological Chemistry 236, 54–60. Griesinger, C., Otting, G., Wuthrich, K. and Ernst, R.R. (1988) Clean TOCSY for 1H spin system identification in macromolecules. Journal of the American Chemical Society 110, 7870–7872. Hansen, C.A., Inubushi, T., Williamson, M.T. and Williamson, J.R. (1989) Partial purification of inositol polyphosphate 1-phosphomonoesterase with characterization of its substrates and products by nuclear magnetic resonance spectroscopy. Biochimica et Biophysica Acta 1001, 134–144. Irvine, R.F. (1986) The structure, metabolism, and analysis of inositol lipids and inositol phosphates. In: Putney, J.W. (ed.) Phosphoinositide and Receptor Mechanisms. Alan R. Liss, New York, pp. 89–107. Isbrandt, L.R. and Oertei, R.P. (1980) Conformational states of myo-inositol hexakis(phosphate) in aqueous solution. A 13C NMR, 31P NMR, and Raman spectroscopic investigation. Journal of the American Chemical Society 102, 3144–3148. IUB Nomenclature Committee (1989) Numbering of atoms in myo-inositol. Biochemical Journal 258, 1–2. IUPAC Commission on the Nomenclature of Organic Chemistry and IUPAC–IUB Commission on Biochemical Nomenclature (CBN) (1976) Nomenclature of cyclitols. Biochemical Journal 153, 23–31. Johnson, K., Barrientos, L.G., Le, L. and Murthy, P.P.N. (1995) Application of 2D TOCSY for structure determination of individual inositol phosphates in a mixture. Analytical Biochemistry 231, 421–431. Johnson, L.F. and Tate, M.E. (1969) Structure of ‘phytic acids’. Canadian Journal of Chemistry 47, 63–73. Lasztity, R. and Lasztity, L. (1990) Phytic acid in cereal technology. Advances in Cereal Science and Technology 10, 309–371. Lemieux, R.U., Kullnig, R.K., Bernstein, H.J. and Schneider, W.G. (1957) Configurational effects in the proton magnetic resonance spectra of acetylated carbohydrates. Journal of the American Chemical Society 80, 1005–1006. Lemieux, R.U., Kullnig, R.K., Bernstein, H.J. and Schneider, W.G. (1958) Configurational effects on the proton magnetic resonance spectra of six-membered ring compounds. Journal of the American Chemical Society 80, 6098–6105. Lindon, J.C., Baker, D. J., Farrant, R.D. and Williams, J.M. (1986) 1H, 13C, and 31P NMR spectra and molecular conformation of myo-inositol 1,4,5-trisphosphate. Biochemical Journal 233, 275–277. Mayr, G.W. and Dietrich, W. (1987) The only inositol tetrakisphosphate detectable in avian erythrocytes is the isomer lacking phosphate at the position 3: an NMR study. Federation of European Biochemical Societies Letters 213, 278–282. Murthy, P.P.N. (2006) Structure and nomenclature of inositol phosphates, phosphoinositides, and glycosylphosphatidylinositols. In: Lahiri Majumdar, A. and Biswas, B.B. (ed.) Biology of Inositols and Phosphoinositides. Springer-Verlag, Berlin, pp. 1–20. Parthasarathy, R. and Eisenberg, F. Jr (1986) The inositol phospholipids: a stereochemical view of biological activity. Biochemical Journal 235, 313–322. Parthasarathy, R. and Eisenberg, F. Jr (1990) Biochemistry, stereochemistry, and nomenclature of the inositol phosphates. In: Reitz, A.B. (ed.) Inositol Phosphates and Derivatives: Synthesis, Biochemistry, and Therapeutic Potential. American Chemical Society, Washington, DC, pp. 1–19. Paton, G., Noailly, M. and Mossoyan, J.C. (1999) Conformational preferences and intramolecular interactions on myo-inositol hexakisphosphoric acid by 1H and 31P NMR studies. Journal of Physical Organic Chemistry 12, 101–107. Posternak, T. (1965) The Cyclitols. Holden-Day, San Francisco, California, pp. 7–49. Raboy, V., Gerbasi, P., Young, K.A., Stoneberg, S., Pickett, S.G., Bauman, A.T., Murthy, P.P.N. and Sheridan, W.F. (2000) Origin and seed phenotype of maize low phytic acid 1-1 and low phytic acid 2-1. Plant Physiology 124, 355–368.

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Schlewer, G., Guedat, P., Ballereau, L., Schmitt, L. and Spiess, B. (1999) Inositol phosphates: infrastructure physico-chemical studies: correlation with binding properties. In: Bruzik, K.S. (ed.) Phosphoinositides: Chemistry, Biochemistry and Biomedical Applications. American Chemical Society, Washington, DC, pp. 255–270. Schmitt, L., Bortmann, P., Schlewer, G. and Spiess, B. (1993) myo-Inositol 2,4,5-triphosphate and related compounds’ protonation sequence: potentiometric and 31P NMR studies. Journal of the Chemical Society, Perkin Transactions 2, 2257–2263. Szwergold, B.S., Graham, R.A. and Brown, T.R. (1987) Observation of inositol pentakis- and hexakisphosphates in mammalian tissue by 31P NMR. Biochemical and Biophysical Research Communications 149, 874–881. Turner, B.L. and Richardson, A.E. (2004) Identification of scyllo-inositol phosphates in soil by solution phosphorus-31 nuclear magnetic resonance spectroscopy. Soil Science Society of America Journal 68, 802–808. Turner, B.L., Papha´zy, M. J., Haygarth, P.M. and McKelvie, I.D. (2002) Inositol phosphates in the environment. Philosophical Transactions of the Royal Society, London, Series B 357, 449–469. Varian (1998) VNMR Command and Parameter Reference Manual, Version 6.1B. Varian, Palo Alto, California, pp. 532–533 and 556. Volkmann, C. J. (2002) NMR investigations of inositol phosphates. MS thesis, Michigan Technological University, Michigan. Volkmann, C. J., Chateauneuf, G.M., Pradhan, J., Bauman, A.T., Brown, R.E. and Murthy, P.P.N. (2002) Conformational flexibility of inositol phosphates: influence of structural characteristics. Tetrahedron Letters 43, 4853–4856. Yang, P., Murthy, P.P.N. and Brown, R.E. (2005) Synergy of intramolecular hydrogen bonding network in myoinositol 2-monophosphate: theoretical investigations into the electronic structure, proton transfer, and pKa. Journal of the American Chemical Society 127, 15848–15861.

3

High-performance Chromatographic Separations of Inositol Phosphates and Their Detection by Mass Spectrometry William T. Cooper1, Matthew Heerboth1 and Vincent J.M. Salters2 1

Department of Chemistry and Biochemistry, Florida State University, Tallahassee, FL 32306, USA; 2National High Magnetic Field Laboratory and Department of Geological Sciences, Florida State University, Tallahassee, FL 32306, USA

Mass spectrometry (MS) would appear to be an attractive approach for measuring inositol phosphates in soils, waters and plant tissues. These compounds are difficult to detect by conventional ultraviolet–visible spectroscopy because they contain no chromophoric groups. The common approach to quantitatively measuring inositol phosphates is their isolation, oxidation and colorimetric detection. Although this approach is relatively simple and straightforward, it can be tedious, time-consuming and subject to interferences. It is also non-specific, in that all inositol phosphates generate the same signal and they cannot be distinguished by detection alone. We have demonstrated that electrospray ionization (ESI) combined with high- and ultrahigh-resolution MS can be used for qualitative organic phosphorus speciation (Llewelyn et al., 2002), i.e. identification of molecular masses and molecular formulas of individual organic phosphorus compounds. We also observed that the detection of individual organic phosphorus compounds within a complicated background matrix of natural organic matter could be difficult. To overcome this problem we recently turned our attention to coupling liquid chromatography separations with inductively coupled plasma (ICP) and ESI–MS, focusing on the qualitative and quantitative detection of individual inositol phosphates. High-performance size-exclusion

chromatography (HP-SEC) and ion-pairing reversed-phase liquid chromatography (HP-SEC) and reversed-phase high performance liquid chromatography (RP-HPLC) have been evaluated for their ability to separate inositol phosphates based on their degree of phosphorylation and isomeric form. Unfortunately, none of these techniques can be fully optimized for inositol phosphate separations because of the limitations imposed by the ICP and ESI steps required for MS detection. These ionization techniques require low concentrations of organic modifiers and volatile salts in the spray matrices, greatly limiting the separation potential of the liquid chromatography methods. To date, HP-SEC has proven to be the most versatile, though least efficient, separation method. However, we also observed that manipulation of various parameters within the ESI source could yield resolvable signals for all six inositol phosphates, allowing detection even when the chromatographic separation is not entirely satisfactory. HP-SEC chromatography of inositol phosphates with ESI–time-of-flight (TOF)–MS detection will be the primary focus of this chapter. ICP ionization combined with high-resolution elemental MS provides very sensitive phosphorus-specific detection and is ideal for quantifying low levels of organic phosphorus in complex matrices. However, the ESI process, in contrast to ICP

©CAB International 2007. Inositol Phosphates: Linking Agriculture and the Environment (eds B.L. Turner, A.E. Richardson and E.J. Mullaney)

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W.T. Cooper et al.

ionization, largely preserves the molecular integrity of organic phosphates. When combined with a high-resolution mass spectrometer, it is useful for verifying the presence of target analytes and identifying new compounds. For this work we have combined ESI with a TOF mass spectrometer that employs reflectron geometry. This new generation TOF instrument includes the most important features of a mass spectrometer used as a chromatographic detector: speed, high mass resolution and good mass accuracy.

We begin with a brief review of classical methods used for separating inositol phosphates before discussing more modern, high-resolution techniques. The classical methods, although modest in terms of capabilities relative to currently available techniques, nevertheless provided much of the early information on inositol structures and occurrence in plants, soils and natural waters. Discussion of these early techniques in relation to the analysis of environmental samples can be found in a recent review (Turner et al., 2002).

Analytical Separations of Inositol Phosphates

Classical methods

Note that in general we will restrict this discussion to analytical separations, which include those techniques that are primarily designed to maximize the resolution of individual compounds within complex mixtures and quantify them. This is in contrast to preparative separations, which are designed to maximize the recovery and/or purification of a target compound. Today, analytical separations are almost always carried out using gas or liquid chromatography, or the very powerful capillary electrophoresis technique. Even though gas chromatography with open-tubular capillary columns provides the greatest separation capabilities of all these methods, the need to volatilize compounds and move them in the gas phase through a liquidcoated capillary column precludes the routine use of gas chromatography for large, ionic, nonvolatile organic phosphate compounds. High-performance liquid chromatography (HPLC) and capillary electrophoresis are thus the techniques of choice for separating inositol phosphates. Separating positional isomers of inositol phosphates is challenging due to the identical number of phosphate groups present. In addition to optimizing the actual separation, a suitable detection procedure must be selected. Inositol phosphates have no distinct chromophoric groups and absorb ultraviolet and visible light only weakly. Thus, traditional ultraviolet and visible detectors used in liquid chromatography and capillary electrophoresis cannot be used without a post-column derivatization step. Sophisticated detectors will thus be necessary for widespread analytical separations of inositol phosphates, including those involving MS.

Most of the first truly quantitative separations of inositol phosphates used low-pressure anion exchange chromatography. Cosgrove (1963, 1966, 1969) reported procedures by which a few inositol phosphate species could be separated. These invariably required harsh solvent conditions (e.g. 0.50 M HCl) and resolved only a few target inositols. McKercher and Anderson (1968a,b) also demonstrated the utility of anion exchange columns for inositol phosphate separations. However, inositol phosphates cannot be reliably quantified by ion exchange alone, due to the co-elution of other organic phosphates. This can be overcome by using hypobromite oxidation prior to chromatography to oxidize interfering substances without degrading the higher-order inositol phosphates (Irving and Cosgrove, 1981). Paper chromatography (Cosgrove, 1980) and gas–liquid chromatography (Irving and Cosgrove, 1982) have also been used to investigate inositol phosphates in environmental samples. The latter technique requires derivatization of the polar, ionized phosphate groups, a step which adds complexity and compromises analytical precision. Paper chromatography can yield fine resolution, but quantification is difficult. However, the technique proved useful for preparative separations that preceded a series of MS (L’Annunziata and Fuller, 1976), infra-red spectroscopy (L’Annunziata et al., 1977) and proton nuclear magnetic resonance (NMR) spectroscopy (L’Annunziata and Fuller, 1971) experiments on inositol phosphates extracted from leaf litter and forest soils. The microbially mediated epimerization of radioactive 14 C(U)-myo-inositol phosphate was even followed by direct autoradiography of paper chromatogra-

HPC Separations and MS Detection

phy bands (L’Annunziata et al., 1977). These experiments, which were highly sophisticated at the time, are summarized in detail elsewhere (L’Annunziata, Chapter 4, this volume). One separation scheme of particular interest used cross-linked dextran gels to separate organic and inorganic phosphorus using Sephadex G-25, with further separation of the organic phosphorus fraction using Sephadex G-50 (Steward and Tate, 1971). Although individual organic phosphorus compounds such as inositols could not be resolved, these latter experiments provided an estimate of the relative size of organic phosphorus pools after hydrolysis with HCl.

Reversed-phase high-performance liquid chromatography The most straightforward liquid chromatography technique is reversed-phase high-performance liquid chromatography (RP-HPLC). Note that high performance refers to liquid chromatography columns packed with small (<10 µm diameter) silica gel particles that normally have an active organic liquid chemically bonded to their surface. When the liquid is hydrophobic (e.g. octane (C8) or octadecane (C18)), the column is used with a polar mobile phase to produce reversed-phase separations based on solute hydrophobicity. The more hydrophobic the solute, the longer is its retention time. RP-HPLC separation usually requires a simple mobile phase comprising water and a polar organic solvent such as methanol, which means that they are ideal when sensitive detectors or post-column derivatization reactions are used. Separation of inositol phosphates based on the degree of phosphorylation using RP-HPLC is a relatively useful technique, as adding –PO3 groups changes the hydrophobicity of the inositol phosphates dramatically. Harland et al. (2004) quantified myo-inositol hexakisphosphate in snack foods with a C18 column and photodiode array multiple wavelength detector after post-column derivatization with Wade’s reagent. Kasim and Edwards (1998) developed an RP-HPLC system that separated myo-inositol trisphosphate through myo-inositol hexakisphosphate, as well as positional isomers of the tetra- and pentakisphosphates, in animal feeds. Casals et al. (2002)

25

quantified higher-order inositol phosphates using a similar system that included post-column derivatization with yttrium, which reduced the absorbance of an organic–yttrium complex in the presence of inositol phosphates. Figure 3.1a is a schematic representation of their post-column reactor and is typical of such reactors, which are designed with small volumes to prevent degradation of chromatographic efficiency. Figure 3.1b is a representative chromatogram demonstrating that straightforward RP-HPLC can, when effectively optimized, separate multiple inositol phosphates, including positional isomers. Note, however, the rather lengthy time required to complete the chromatogram.

Ion-pairing reversed-phase high-performance liquid chromatography Strongly acidic buffers were required in both the RP-HPLC separations of inositol phosphates described above because inositol phosphates in anionic form are hydrophilic and do not partition into the non-polar stationary phase (C18 in the references cited) to any extent. Buffers are thus used to suppress ionization and increase the affinity of inositol phosphates for the non-polar stationary phase. An alternative to this ionization suppression approach is to put hydrophobic cations that form ion pairs with phosphate in the mobile phase. By adding such an ion-pairing reagent (e.g. tetrabutylammonium chloride) to the mobile phase, the normally hydrophobic stationary phase becomes coated with the relatively hydrophobic cation (tetrabutylammonium). Phosphate anions can then form ion pairs with the exposed positive charges on the stationary phase surface, providing an ion-exchange-type mechanism. This technique is referred to as ionpairing RP-HPLC (e.g. Brando et al., 1990). The potential of such separations coupled to MS detection is demonstrated in Fig. 3.2a, in which a mixture of organic phosphate standards was resolved in <20 min (Cooper et al., 2005a). This system was used to qualitatively identify organic phosphorus compounds in a treatment wetland (Fig. 3.2b). This approach holds great promise for analysis of inositol phosphates because the ionexchange mechanism appears to provide the best separations of these compounds.

26

W.T. Cooper et al.

100

Ins(1,3,4,6)P4

40

Standard connector

20 Input PAR solution

Input eluent Magnetic bar

Ins(3,4,5,6)P4

mV

60

InsP6

80

Ins(1,3,4,5,6)P5

Output

Inositol polyphosphates detected in three pigeon erythrocyte samples Mean ⫾ SD (µmol/ml erythrocyte) Ins(1,3,4,6)P4 <0.005 Ins(3,4,5,6)P4 0.009⫾0.005 Ins(1,3,4,5,6)P5 3.190⫾0.34 0.052⫾0.019 InsP6

Stirrer 0 20

(a)

(b)

30

40

50

Elution time (min)

Fig. 3.1. (a) Post-column reactor for detecting non-absorbing inositols with metal dye reaction. (b) Reversed-phase high-performance liquid chromatography (RP-HPLC) separation of minor inositol phosphates in blood cells. (From Casals et al., 2002. Reprinted with permission from Elsevier Science.)

Ion exchange and ion chromatography Currently, ion-exchange chromatography with strong anion exchange columns is the most suitable separation technique for inositol phosphates. The ion-exchange mechanism relies on a specific anion (phosphate)–cation (column) interaction. Increasing the number of these interactions by adding phosphate groups increases retention dramatically. Further, the strength of a phosphate–cationic site interaction includes a significant steric component, and thus ion exchange has proven effective in separating inositol phosphates and their positional isomers. Skoglund et al. (1998) separated 25 distinct inositol phosphates using two strong anion exchange columns followed by post-column derivatization and ultraviolet detection. Ion chromatography is even more powerful for inositol phosphate separations because in principle it allows sensitive conductivity detection. The technique combines an ion-exchange separation column (anion or cation) with a second suppressor column, which reduces the background conduc-

tivity of the mobile phase and allows conductivity detection of any analyte ion. Ion chromatography thus satisfies both requirements for analytical separations of inositol phosphates: selective retention mechanisms that allow chromatographic separation of virtually all target analytes, plus sensitive detection without post-column derivatization. Phillippy and Johnston (1985) were the first to apply ion chromatography to inositol phosphate analysis, measuring myo-inositol hexakisphosphate in foodstuffs, although they found post-column derivatization and ultraviolet detection more sensitive than conductivity. A number of research groups subsequently used ion chromatography to detect inositol phosphates. Figure 3.3 demonstrates the application of such a system for studying the enzymatic dephosphorylation of myo-inositol hexakisphosphate, in which 27 inositol phosphates were separated and detected (Chen and Li, 2003). This particular separation included a gradient elution that allowed weakly retained inositol phosphates to be separated in a weak mobile phase, and then more strongly retained inositol phosphates to be eluted by the increasingly stronger mobile phase.

HPC Separations and MS Detection

120,000

1− 2− 3,4− 5− 6,7− 8−

6

100,000

1

CPS (P)

80,000 3 60,000 2

7

4

20,000

Capillary electrophoresis

8

5 40,000

o-P AMP CP ADP PEP ATP

0 0

(a)

5

10

15 20 Time (min)

25

30

o-P

12,000

CPS (P)

10,000 8,000 6,000 4,000 2,000 0

0

5,000

5

10

15 20 25 Time (min)

30

23.4

o-P

35

40

36.1

CPS (P)

4,000 3,000 2,000 1,000 0

(b)

0

5

10

15 20 Time (min)

25

30

35

27

40

Fig. 3.2. (a) Optimized separation of organic phosphate standards by ion-pair chromatography with direct, on-line phosphorus-specific inductively coupled plasma mass spectrometry (ICP–MS) detection. AMP, ADP, ATP = adenosine mono-, diand triphosphate; CP = creatine phosphate; PEP = phosphoenolpyruvate. The peak at ~5 min is phosphate. (b) Ion-pair high-performance liquid chromatography (ion-pair HPLC) separation of organic phosphorus compounds with phosphorusspecific ICP–MS detection from the treatment wetland inflow (top) and outflow (bottom). CPS = counts per second. (Reprinted from Cooper et al., 2005b.)

The chromatogram in Fig. 3.3 demonstrates both the advantages and disadvantages of gradient elution liquid chromatography, because although the 27 inositol phosphates were well resolved in <50 min, significant drift in the detector baseline signal occurred as the composition of the mobile phase changed.

Any review of separations of inositol phosphates must include a discussion of capillary electrophoresis, a very powerful analytical separation technique. It is not a chromatographic technique, because it separates solutes based on electrophoretic mobility rather than affinity for a stationary phase. Sample mixtures are loaded into a glass capillary tube filled with electrolyte solution, and a voltage is then applied between the injector and detector ends of the tube. Modern capillary electrophoresis evolved from classical electrophoresis in the early 1980s when it was recognized that the use of small-diameter glass tubes (i.e. capillary tubes of 0.25–0.50 mm i.d.) provided efficient heat transfer and minimized the effects of Joule heating. This meant that high voltages (10–30 kV) could be applied over fairly lengthy distances, resulting in efficient separation of solutes with differing electrophoretic mobility. Capillary electrophoresis separation of inositol phosphates appears to be straightforward, as addition of –PO3 groups to the inositol ring should change electrophoretic mobility dramatically. In addition, resolution of positional isomers can often be accomplished by adding certain solvents to the electrolyte solution (e.g. cyclodextrins). However, only a few reports have appeared in the literature describing capillary electrophoresis separations of inositol phosphates. Henshall et al. (1992) separated myo-inositol 1,4-bisphosphate, myo-inositol 1-monophosphate, and a mixture of myo-inositol 1,4,5-trisphosphate and myo-inositol hexakisphosphate. They were also able to separate DL-myoinositol 1-monophosphate and DL-myo-inositol 2-monophosphate with a second electrolyte solution. Buscher et al. (1994) used ‘indirect detection’ to identify all six inositol phosphate esters by capillary electrophoresis, although the resolution of myo-inositol penta- and hexakisphosphates was poor (Fig. 3.4). Taguchi et al. (2000) took a different approach, using capillary electrophoresis to separate phospholipids into broad classes, followed by ESI–MS detection. Rather than physically separating the components of each class, they relied on in-source fragmentation to generate mass spectral patterns that could be used to identify several components within each class. The principal drawback to capillary electrophoresis as a separation technique for inositol

28

W.T. Cooper et al.

27

0.5 0.06 2

0.4

25

24

3 4 5

1

6

26

23

CPS

0.3 0.01 4.9

0.2

5.4

1617 19 20 21 18

5.9 14

22

15

11

0.16

0.1

10 9

0.13 0

0.10

78

12 13

0.07 12.5 13.5 14.5 15.5 16.5

−0.1 10.0

0

20.0

30.0

50.0

40.0

Time (min) Fig. 3.3. Separation of 27 inositol phosphate standards by ion chromatography. (From Chen and Li, 2003. Reprinted with permission from Elsevier Science.) See citation for identification of inositol phosphates and chromatographic conditions. CPS = counts per second.

phosphates is the low ultraviolet and visible light absorbances of the target compounds. Post-column derivatization, which is often used in HPLC separations, is not possible in capillary electrophoresis, so detection normally requires a ‘bulk solution’ effect such as decrease in absorp-

tion of a chromophore added to the electrolyte solution. From our experience, this complicates optimization of experimental parameters such as applied voltage, injection time and electrolyte composition, and even optimized capillary electrophoresis systems are far from robust.

4

0.02

UV absorbance

2 1

0.01

3

5 6

0

−0.01

−0.02 0.00

1.00

2.00

3.00

4.00

5.00

6.00

Time (min) Fig. 3.4. Capillary-zone electrophoresis separation of inositol phosphates with indirect ultraviolet detection at 214 nm. (From Buscher et al., 1995. Reprinted with permission from Elsevier Science.) 1 = IP6; 2 = IP5; 3 = IP4; 4 = IP3; 5 = IP2; 6 = IP1.

HPC Separations and MS Detection

Furthermore, changes in the electrophoretic mobility of inositol phosphates with the addition of phosphate groups appear to be more complicated than first thought.

Mass Spectrometry Detection There are surprisingly few studies in the literature describing the use of MS for organic phosphorus detection in natural systems, and even fewer describing the measurement of inositol phosphates. Several studies attempted to detect low levels of inositol phosphates in biochemical matrices such as red blood cells (Sherman et al., 1986; Hsu et al., 1990, Buscher et al., 1995), but were largely unsuccessful until the introduction of ESI (Fenn et al., 1989). Organic phosphorus is dominated by the PO4 group(s) that impart(s) polarity and mass. It is thus difficult to ionize organic phosphorus molecules without substantial fragmentation, complicating the qualitative identification of individual compounds. It was not until the advent of soft ionization techniques, primarily ESI and matrix-assisted laser desorption ionization, that MS of molecules such as proteins, peptides and inositol phosphates became possible. Kerwin et al. (1994) described the role of glycerolphosphoinositols in cell membranes using both positive and negative ionmode ESI–MS. However, it is now apparent that the negative ion mode is superior (Hsu and Turk, 2000).

Electrostatic lenses Quadrupole mass spectrometer

Cylindrical electrode

29

Electrospray ionization The importance of the ionization step in MS detection of inositol phosphates, as well as the limitations it imposes on the chemical nature of the solvent in which inositol phosphates are dissolved, means that a brief discussion of the most important ESI process for organic phosphorus MS is necessary. Figure 3.5a is a schematic representation of an original ESI source design. A liquid solution containing the analyte(s) is pumped through a steel capillary tube held at a potential of 1–3 kV relative to a counter electrode in the source. The electrostatic field generated by the applied potential disperses the emerging solution into a fine mist of charged droplets. These droplets are rapidly desolvated, leaving charged analytes that are then introduced into the mass spectrometer through a ‘skimmer cone’, which has the same charge as the gas-phase ions. The voltage of this cone is important, as it needs to be high enough to focus and guide sufficient ions into the mass analyser to produce a signal, but at the same time low enough so that it does not induce ‘insource fragmentation’. Both positive and negative ions can be measured in this way, depending on the polarity of the capillary tip and cone. ESI mass spectra are normally dominated by intact molecular ions, often with multiple charges. Solvent composition is critical for efficient ionization by the electrospray method. The Liquid chromatography eluent

Nebulizer gas Desolvating chamber

Capillary Needle

Orifice 2

Liquid sample

Ion guide

Desolvating gas Skimmer Ring lens Drying gas

(a)

First pumping stage

Second pumping stage

Orifice 1

(b)

Fig. 3.5. Schematic representation of electrospray ionization (ESI) sources: (a) The original design of Fenn et al. (1989); (b) orthogonal design used in the JEOL AccuTOF mass spectrometer. (Courtesy of JEOL, Inc., Tokyo, Japan.)

30

W.T. Cooper et al.

solvent must support and encourage ionization of the analyte(s), but it cannot be better than the analyte in taking up the charge applied by the applied field. Thus, when forming negative analyte ions (e.g. phosphate ions), very weak bases must be used for buffering. For example, we typically use ammonium bicarbonate, which is both a weak buffer and quite volatile. Small amounts of methanol or acetonitrile are also commonly added to buffered aqueous solutions to reduce the surface tension of droplets. These constraints make the mobile phases used in strong anion exchange chromatography unacceptable for ESI–MS detection, and have been one driving force behind the interest in finding alternative separation schemes for inositol phosphates. Dragani et al. (2004) demonstrated a promising approach for analysing glycerophosphoinositol using a β-cyclodextrin bonded phase RP-HPLC system with ESI–MS detection.

Time-of-flight mass analysis MS produces information about the masses and abundances of gas-phase ions. It is a very powerful analytical technique for determining the elemental composition and chemical formulas of molecules, as molecular mass is the single most important piece of information necessary for characterizing chemical structures. MS is also an important quantitative tool because mass spectrometers are in general very sensitive and will respond to most compounds in a predictable way over large concentration ranges. It should be noted that in the MS community the currently accepted unit for atomic or molecular mass is the dalton (Da), which is exactly one-twelfth the mass of a 12C atom. A mass spectrum is thus a plot of ion intensity as a function of mass/charge ratio (m/z). To get the mass of an ion its charge must be known, but this can usually be determined from the spectrum. Many molecules will fragment during the ionization step, and their spectra then comprise a molecular ion and a series of fragment ions. Although such fragmentation will produce complicated spectra if many compounds are present, it nevertheless can provide valuable information

about the sub-structures present in the original molecules. TOF mass analysers are velocity spectrometers, in that ions are separated by m/z based upon different velocities acquired in an electrostatic field. As all ions are accelerated through the same field, they all have the same nominal kinetic energy. Thus, their velocities will vary inversely with m/z. A mass spectrum is obtained by monitoring ions arriving at the detector as a function of time. The defining relationship that equates drift time to m/z is given in 1/2

L 1 m t= v =Ld 2V nd z n

(3.1)

where t is the time for the ion to reach the detector, L the length of the tube, v the ion velocity and V the accelerating voltage. Early TOF analysers did not have particularly good resolution, generally less than about 500, but their speed and virtually unlimited mass range made them popular. However, great improvements in the resolution of TOF spectrometers followed from introduction of the reflectron (Fig. 3.6), which incorporates a series of electrical lenses that compensate for variations in ion velocity (Cooper et al., 2005b). Modern reflectron TOF analysers are fast, which makes them ideal for use as chromatographic detectors, and also have high mass resolving power and mass accuracy, a necessary requirement for detecting molecules like inositol phosphates in complex matrices.

Field-free drift region

Reflectron lenses

Detector Ion-storage region Deflection plates

Fig. 3.6. Schematic representation of a time-offlight (TOF) mass analyser with reflectron geometry. Ions that enter the field-free drift region migrate to the detector at a rate that is dependent on their mass/charge ratio (m/z). The reflectron lenses compensate for variations in kinetic energies of the inject ions; these variations would otherwise produce broadened peaks and loss of spectral resolution.

HPC Separations and MS Detection

Electrospray Ionization Molecular Mass Spectrometry of Inositol Phosphates Direct electrospray ionization time-offlight mass spectrometry of inositol phosphates In previous studies of dissolved organic phosphorus speciation in the Florida Everglades we noted that ESI was inefficient for organic phosphorus (Llewelyn et al., 2002). Figure 3.7 includes ultrahigh-resolution ESI mass spectra of organic phosphorus standards before and

31

after the standards had been concentrated by a phosphorus-specific isolation procedure. The mass spectrum of the mixture before precipitation shows that many of these standards were not seen as simple [M + H]+ ions. In addition, ESI efficiencies of organophosphates do not appear to be particularly high. These observations further reinforce the need for selective concentration of dissolved organic phosphorus. Fortunately, inspection of Fig. 3.7b suggests that the organic phosphorus isolation procedure we use does not alter the ESI characteristics of these compounds. We carried out a series of initial experiments using a ‘standard’ skimmer voltage setting

[AMP + H]+

[PEP + Na]+

[AMP + Na]+ [TP +

(a)

[2 RP + H]+

H]+

250

300

350

400 m/z

450

500

550

[PEP + Na]+ [AMP + H]+

[AMP + Na]+

[2 RP + H]+

[TP + H]+

250

300

350

400

450

500

550

(b) Fig. 3.7. Positive ion ESI 9.4 T Fourier-transform ion-cyclotron resonance mass spectra showing the standard dissolved organic phosphorus mixture – tyrosine phosphate (TP), adenosine monophosphate (AMP), phosphoenolpyruvate (PEP) and ribose phosphate (RP) – (a) prior to barium precipitation and (b) following barium precipitation. (From Llewelyn et al., 2002. Reprinted with permission from the American Chemical Society.)

32

W.T. Cooper et al.

Table 3.1. Isomeric forms of the six inositol phosphates used in this work, with molecular weights and m/z value at which the molecular ion of each appears in the electrospray ionization–time-of-flight (ESI–TOF) mass spectrum.

Inositol phosphate

Isomer

IP1 IP2 IP3 IP4 IP5 IP6

myo-Inositol myo-Inositol myo-Inositol myo-Inositol myo-Inositol myo-Inositol

2-monophosphate 2,4-bisphosphate 1,4,5-trisphosphate 1,4,5,6-tetrakisphosphate 1,3,4,5,6-pentakisphosphate hexakisphosphate

Molecular weight (Da)

Molecular ion peak (Da)

260.0 340.0 420.0 499.9 579.9 659.9

259.0 339.0 419.0 498.9 578.9 658.9

observe what appeared to be significant fragmentation of myo-inositol hexakisphosphate, with the bis- and pentakisphosphates also appearing in the mass spectrum (Fig. 3.8). Although the presence of lower-order inositol phosphates as impurities in the myo-inositol hexakisphosphate standard cannot be ruled out, we believe that the phosphate groups in inositol phosphates may be relatively labile during ESI, as the mass differences we observe here (80 Da) represent the loss of HPO3 group(s). This fragmentation process has been observed before with phosphate-containing peptides (Neubauer and Mann, 1999). Thus, intact molecular ions may not always be the most abundant in the mass spectra of inositol phosphates. Nevertheless, these results are encouraging because they indicate that with sufficiently good analytical separation the molecular formulas of

of 60 V. A JEOL AccuTOF mass spectrometer ( JEOL, Inc., Tokyo, Japan) was used for all the MS measurements described here. This is a single-stage reflectron mass analyser with a mass resolution of ~6000 and mass accuracy of less than 5 ppm. This instrument includes an ESI source with orthogonal geometry (Fig. 3.5b). Inositol phosphates were obtained from Sigma Chemicals (St Louis, Missouri, USA). Table 3.1 summarizes the inositol phosphates used in these studies, as well as molecular weights of the intact inositol phosphate and its molecular ion m/z value (M – H). In the first experiment a mixture of sodium salts of myo-inositol monophosphate (C6H6 (HnPO4)1; Mw = 260 Da) and myo-inositol hexakisphosphate (C6H6(HnPO4)6; Mw = 660 Da) was efficiently ionized, but we were surprised to

419 Da

259 Da

IP6; 659 Da

339 Da 498 Da

579 Da

100

200

300

400 m/z

500

600

700

800

Fig. 3.8. Electrospray ionization–time-of-flight (ESI–TOF) mass spectrum of myo-inositol hexakisphosphate (IP6); skimmer cone voltage 60 V.

HPC Separations and MS Detection

the individual inositol phosphate species can be verified by ESI–MS. In addition, these results suggest that it may be possible to generate a common ion from different inositol phosphates using in-source fragmentation. This would allow phosphorus-specific detection of peaks emerging from HPLC separation columns. These initial experiments were followed by a study to better understand the stability of inositol phosphates in the ESI source. This same mixture of myo-inositol monophosphate and myoinositol hexakisphosphate was sprayed under identical conditions, but varying skimmer voltages. The dramatic effect of skimmer voltage is evident in the mass spectra of Fig. 3.9. At 20 V (Fig. 3.9a) myo-inositol hexakisphosphate at 659 Da is barely visible, while myo-inositol monophosphate at 259 Da is quite intense. However, increasing the skimmer voltage to 80 V (Fig. 3.9b) greatly increases the myo-inositol hexakisphosphate molecular ion peak at 659 Da, while the myo-inositol monophosphate peak is not as intense as at 20 V. Figure 3.10 summarizes molecular ion peak intensities over a wide range of skimmer voltages. Clearly, there is an optimum skimmer voltage between 40 and 80 V at which both molecular ion peaks are intense and suitable for use in selected ion monitoring (SIM) to detect specific inositol phosphates. This conclusion is confirmed by the mass spectrum of a mixture of six inositol phosphates (Fig. 3.11) in

33

which the molecular ions (as M – H) of each compound are visible. The low intensities of molecular ions at high skimmer voltages summarized in the data of Fig. 3.10 also suggest that it might be possible to identify a fragmentation product common to all the inositol phosphates and exploit in-source fragmentation for SIM after chromatographic separation. One potential fragment appears in the spectra of both myo-inositol monophosphate and myo-inositol hexakisphosphate at a skimmer voltage of 100 V at 79 Da and corresponds to PO3. This prominent fragment was previously observed by Neubauer and Mann (1999) in a study of phosphopeptide phosphorylation using triple quadrupole MS. We therefore carried out another series of experiments in which we monitored the intensities of molecular ions of all six inositol phosphates and this 79 Da fragment at skimmer cone voltages of 20 and 60 V. Intensity ratios of molecular ion/fragment ion for the inositol phosphates are plotted in Fig. 3.12. From these data it is clear that all inositol phosphates fragment to a significant extent at 60 V. However, there is a very interesting trend at 20 V. The tendency to fragment, or at least to lose PO3, appears to increase as HPO3 groups are added to the inositol backbone. These results have somewhat negative connotations regarding the analysis of inositol phosphate

IP1; 259 Da

IP1; 259 Da IP6; 659 Da

IP6; 659 Da

200 (a)

400

600 m/z

800

200 (b)

400

600 m/z

Fig. 3.9. ESI–TOF mass spectra of a myo-inositol monophosphate (IP1) and myo-inositol hexakisphosphate (IP6) mixture at skimmer cone voltages of (a) 20 V and (b) 80 V.

800

34

W.T. Cooper et al.

25,000

Intensity

20,000

IP1 IP6

15,000 10,000 5000 0 20

40

60

80

100

120

140

Cone voltage (V) Fig. 3.10. Molecular ion peak intensities of myo-inositol monophosphate (IP1) and myo-inositol hexakisphosphate (IP6) as a function of skimmer cone voltage.

mixtures by direct ESI–MS. ESI of any inositol phosphate mixture will apparently produce molecular ions of each compound, but fragment ions will appear at the same m/z values as less phosphorylated inositols. These results thus reinforce the need for a moderately good analytical separation before the ESI–MS detection.

Size-exclusion chromatography with mass spectrometry detection of inositol phosphates Analysis of inositol phosphates by ESI–TOF–MS appears promising given the results described above. However, when attempting to detect inos-

itol phosphates in complex matrices such as soils and animal manures it is necessary to isolate them from other species that interfere with their ionization in the ESI source. This was one of the primary findings of our work with organic phosphates in an Everglades treatment wetland (Llewelyn et al., 2002): organic phosphorus does not compete very effectively for charge when in a high background of natural organic matter. We thus began an evaluation of HP-SEC as a preliminary, on-line isolation step prior to ESI–TOF–MS of inositol phosphates. SEC would appear to be the most useful liquid chromatography separation technique for this purpose, as the mobile phases normally required are aqueous-based solutions with small amounts of organic modifier(s) and volatile buffer salts. Size-exclusion chromatography

IP6; 659 Da

IP1; 259 Da

IP3; 419 Da

IP4; 498 Da IP5; 579 Da

IP2; 339 Da

200

600

400

800

m/z Fig. 3.11. ESI–TOF mass spectrum of an inositol phosphate mixture at a skimmer cone voltage of 60 V.

SEC is based on a relatively simple principle: larger molecules are ‘excluded’ to a greater extent from the inner spaces of a porous column packing material than smaller molecules, which can penetrate the openings of the small pores. The average residence time within the column thus depends on the ‘effective’ molecular size, as molecules in the pore volume of the packing are removed form the flowing mobile-phase stream and do not move towards the end of the column. Retention in HP-SEC can be described by Vr = Vvoid + KVpores

(3.2)

where Vr is the retention volume of the solute (retention time × volumetric mobile-phase flow

HPC Separations and MS Detection

35

Molecular/fragment ion ratio at 60 V Molecular/fragment ion ratio at 20 V Intensity ratio

4

Intensity ratio

50 40 30

3.5 3 2.5 2 1.5 1 0.5 0

IP3

IP4

IP5

IP6

IP5

IP6

20 10 0

IP1

IP2

IP3

IP4

Fig. 3.12. Ratios of intensities of inositol phosphate molecular ions to fragment ion peak at 79 Da and skimmer cone voltages of 20 and 60 V.

rate), Vvoid is the volume of interparticle void space in the column, Vpores is the total pore volume accessible to the smallest solutes and K is a constant that describes the probability of the solute penetrating into the pore space. The value of K is a complex function of primarily molecular size, but also factors such as shape and chargedensity. Any molecule that is too large to enter any of the pores is totally excluded (K = 0), while a small molecule can penetrate all the pores and has access to the entire pore volume (K = 1). Sizeexclusion columns are thus characterized by an exclusion limit, which is the molecular weight of all molecules with Vr = Vvoid , and total permeation limit, which is the molecular weight of all molecules with Vr = Vvoid + Vpores. Molecules that are of sizes that fall within exclusion and total permeation limits will be separated. The separations described in this chapter were carried out on a Beckman–Coulter System Gold liquid chromatograph. Mobile phases were composed of 4:1 (v/v) mixtures of water/methanol. The water component of the mobile phase was buffered with ammonium bicarbonate at concentrations of 0.10 and 0.01 M. We used a 250 × 4.6 cm i.d. PL-Aquagel-OH polymer column (Polymer Laboratories, Shropshire, UK), 8 µm particle diameter, with a molecular weight separation range of 100–30,000 Da. The column was calibrated with poly(styrene)sulphonate (PSS)

standards with nominal molecular weights of 1640, 7900, 16,600 and 70,000 Da. Such standards are often used to calibrate SEC columns for natural organic matter separations, as they are thought to behave much like natural organic matter. The chromatograms were characteristic of most HPSEC separations: peaks are relatively broad and chromatographic resolution is not great, but molecules that are very different in size can be resolved. SEC was performed using two mobile phases that differed only in their ionic strength. The first consisted of a mixture that was 80% water containing 0.10 M ammonium bicarbonate and 20% methanol. The second mobile phase was identical, except that the aqueous phase was only 0.01 M in ammonium bicarbonate. Previous experiments by Reemtsma and These (2003) demonstrated that separation of natural organic matter improved as the ionic strength of the buffer increased, probably due to ionic interactions between charged solutes and polar sites within the Aquagel polymer matrix. Calibration curves for the two mobile phases are included in Fig. 3.13. These curves demonstrate the classical log-linear relationship between molecular weight and retention time in SEC. They also indicate that much better resolution can be obtained with the higher ionic strength mobile phase. Our results confirm that high ionic strength buffers are necessary for efficient separation of

36

W.T. Cooper et al.

5

log Mw

4.5 (a) (b)

4

3.5

3 15

17

19

21

23

25

Tr (min) Fig. 3.13. Molecular weight (Mw) vs. retention time (Tr) calibration curves for poly(styrene)sulphonate (PSS) standards on a PL-Aquagel-OH polymer column with 8 µm particle diameter: (a) 80% water containing 0.10 M ammonium bicarbonate and 20% methanol; (b) 80% water containing 0.01 M ammonium bicarbonate and 20% methanol.

natural organic matter in size-exclusion polymer columns. However, this presents a dilemma when ESI–MS will be used for detection, since high ionic strength decreases signal intensity. Indeed, baselines in chromatograms using the 0.10 M buffer and ESI–TOF–MS detection were too noisy to be useful. Thus, we were forced to trade the chromatographic resolution obtainable at high buffer strengths with acceptable signal intensities obtainable only at lower buffer strengths. High-performance size-exclusion chromatography with ESI–TOF selected ion monitoring mass spectrometry detection of inositol phosphates The behaviour of inositol phosphates in an ESI source provides a number of opportunities for determining them in complex matrices. As noted previously, the SEC separation nicely isolates the inositol phosphates from much of the interfering natural organic matter that would otherwise obscure their ESI–MS signals due to ionization suppression. Although some compromise between separation efficiency and signal intensity is necessary, acceptable isolation of inositol phosphates can be achieved at lower mobile-phase ionic strengths. Because inositol phosphates ionize at lower cone voltages primarily as molecular ions, individual inositol phosphates can be identified as they elute from an SEC column by monitoring

the appropriate ion as a function of time. This approach is generally referred to as SIM and is now a well-established analytical technique. Our approach to SIM is to scan the entire mass range of interest repeatedly throughout a chromatographic separation. Then, a ‘mass chromatogram’ can be reconstructed by plotting the intensity of one molecular ion as a function of time. SIM is demonstrated in Fig. 3.14. The total ion current chromatogram of a sample of myo-inositol bisphosphate shown in Fig. 3.14a was obtained by summing the intensities of all ions within the mass range being monitored, in this case 80–800 Da. No peaks are visible because the IP2 signal is so low that it does not rise above the background signal produced by ionization of molecules in the mobile phase. However, when the signal produced only by the ion at m/z = 339 Da is plotted, the myo-inositol bisphosphate peak (as M – H) is clearly visible and intense relative to the background. Figure 3.14b is a classic example of a ‘mass chromatogram’, in this case of inositol bisphosphate. The previous experiments on direct ESI–TOF–MS indicated that all six inositol phosphates we tested produced sufficiently intense molecular ion peaks (as M – H) to be monitored by this SIM technique. Figures 3.15 and 3.16 include mass chromatograms for two of the six inositol phosphates, separated as a mixture. It should be noted that both mass chromatograms were obtained with only one size-exclusion sepa-

HPC Separations and MS Detection

(a)

(a)

5

10

15

20

Retention time (min)

5 (b)

37

10 15 Retention time (min)

20

100 (b)

200

300

400 500 m/z

600

700

800

Fig. 3.14. High-performance size-exclusion chromatograms (HP-SEC) with ESI–TOF detection of myo-inositol bisphosphate: (a) Total ion chromatogram; (b) selected ion monitoring (SIM) chromatogram at m/z 339.

Fig. 3.15. HP-SEC and mass spectrum of myoinositol monophosphate: (a) SIM chromatogram at m/z 259; (b) mass spectrum.

ration; the mass chromatograms were reconstructed from this single chromatogram by plotting the intensities of the appropriate masses. Also included in these figures are the mass spectra over the entire mass range monitored (60–800 Da) at the elution time of each compound. The relatively low background and lack of many noninositol phosphate peaks in these spectra are no doubt one additional benefit of the HP-SEC separation prior to mass analysis.

elemental MS of phosphorus at m/z 31. This special approach to SIM in which a fragment rather than a molecular ion is monitored during a chromatographic separation is sometimes referred to as ‘mass fragmentometry’, and the resulting chromatogram, a ‘mass fragmentogram’. A ‘mass fragmentogram’ of a mixture of all six inositol phosphates is depicted in Fig. 3.17. This fragmentogram was reconstructed from continued scanning of masses 60–800, and then plotting the intensity of the 79 Da peak as a function of time. Surprisingly, this SEC column, with a nominal separation range of 100–30,000 Da, was able to resolve the inositol monophosphate (peak at 19 min) from the other inositol phosphates. However, myo-inositol bisphosphate through myoinositol hexakisphosphate were not resolved.

High-performance size-exclusion chromatography with ESI–TOF mass fragmentometry detection of inositol phosphates We previously identified a fragment at m/z 79 that was common to all the inositol phosphates. This PO3 fragment could potentially be monitored in the SIM mode and serve as a signal for any inositol phosphate. This would convert the ESI–TOF mass spectrometer into an inositol phosphate–specific detector, and accomplish what we previously achieved using ICP ionization and

Summary ESI–TOF–MS offers several approaches to measuring inositol phosphates in complex environmental, agricultural and biological matrices. All

38

W.T. Cooper et al.

5

(a)

100

200

15 10 Retention time (min)

300

400

(b)

500

600

20

700

800

m/z

Fig. 3.16. HP-SEC and mass spectrum of myoinositol hexakisphosphate: (a) SIM chromatogram at m/z 659; (b) mass spectrum.

six inositol phosphates are sufficiently stable during the ionization process to be observed as (M – H) molecular ions. Unfortunately, they also fragment to some extent, losing PO3 in a process yielding ions that appear at exactly the same m/z values as other less phosphorylated inositols. Thus, a physical separation by some sort of chro-

matography will be necessary before SIM of each inositol phosphate can be quantitative. However, the tendency to fragment can be used as an advantage, as all inositol phosphates appear to produce PO3 as a common fragment ion. SIM of this fragment at 79 Da can thus convert the ESI–TOF mass spectrometer into a phosphorusspecific chromatographic detector, in the same way that inductively coupled plasma mass spectrometry (ICP–MS) monitoring of elemental phosphorus at 31 Da has been used (Cooper et al., 2005a). The combination of liquid chromatography and MS would thus appear to offer a new, more sensitive and selective analytical method for quantitatively identifying inositol phosphates in environmental samples. MS is significantly more sensitive relative to NMR and also minimizes the effect of other organic phosphates on the quantification of individual inositol phosphates. It has been noted that many of the useful ion-exchange separation techniques require pre-treatment of samples by hypobromite oxidation to remove other organic phosphates that co-elute and interfere with inositol phosphate quantification when phosphorus-specific detectors that include post-column derivatization are used (Irving and Cosgrove, 1981). These interfering phosphorus compounds would not be a problem if MS with SIM were used as the chromatographic detector, eliminating the need for a preliminary oxidation step. However, the ESI process that is critical to MS detection is highly sensitive to salts and polar compounds that compete with phosphates for charge in the ESI source. Thus, complex environmental samples will still require extensive preliminary clean-up and isolation procedures to remove inositol phosphates from metals and organics (e.g. high-molecular weight humic acids) to which they are bound before these sensitive and selective HPLC–ESI– MS techniques could be used.

Acknowledgements

10

15

20

Retention time (min)

Fig. 3.17. HP-SEC SIM at m/z 79 (mass fragmentogram) of an inositol phosphate mixture; skimmer cone voltage 60 V.

This work was supported by grants from the South Florida Water Management District and the United States Department of Agriculture (USDA). The assistance of Dr Umesh Goli, Director of the Mass Spectrometry Laboratory at Florida State University Department of Chemistry, is greatly appreciated.

HPC Separations and MS Detection

39

References Brando, C., Hoffman, T. and Bonvini, E. (1990) High-performance liquid chromatographic separation of inositol phosphate isomers employing a reversed-phase column and a micellar mobile phase. Journal of Chromatography B 529, 65–80. Buscher, B.A.P., Irth, H., Andersson, E., Tjaden, U.R. and van der Greef, J. (1994) Determination of inositol phosphates in fermentation broth using capillary zone electrophoresis with indirect UV detection. Journal of Chromatography 678, 145–150. Buscher, B.A.P., Vanderhoeven, R.A.M., Tjaden, U.R., Andersson, E. and Vandergreef, J. (1995) Analysis of inositol phosphates and derivatives using capillary zone electrophoresis mass-spectrometry. Journal of Chromatography A 712, 235–243. Casals, I., Villar, J.L. and Riera-Codina, M. (2002) A straightforward method for analysis of highly phosphorylated inositols in blood cells by high-performance liquid chromatography. Analytical Biochemistry 300, 69–76. Chen, Q.-C. and Li, B.W. (2003) Separation of phytic acid and other related inositol phosphates by high performance ion chromatography and its applications. Journal of Chromatography A 1018, 41–52. Cooper, W.T., Llewelyn, J.M., Bennett, G.L. and Salters, V.J.M. (2005a) Mass spectrometry of natural organic phosphorus. Talanta 66, 348–358. Cooper, W.T., Llewelyn, J.M., Bennett, G.L., Stenson, A.C. and Salters, V.J.M. (2005b) Organic phosphorus speciation in natural waters by mass spectrometry. In: Turner, B.L., Frossard, E. and Baldwin, D.S. (eds) Organic Phosphorus in the Environment. CAB International, Wallingford, UK, pp. 45–74. Cosgrove, D.J. (1963) The chemical nature of soil organic phosphorus. I. Inositol phosphates. Australian Journal of Soil Research 1, 203–214. Cosgrove, D.J. (1966) Detection of isomers of phytic acid in some Scottish and California soils. Soil Science 102, 42–43. Cosgrove, D.J. (1969) Ion-exchange chromatography of inositol polyphosphates. Annals of the New York Academy of Sciences 165, 677–686. Cosgrove, D.J. (1980) Inositol Phosphates: Their Chemistry, Biochemistry and Physiology. Elsevier, Amsterdam, The Netherlands. Dragani, L.K., Berrie, C.P., Corda, D. and Rotilio, D. (2004) Analysis of glycerolphosphoinositol by liquid chromatography–electrospray ionisation tandem mass spectrometry using β-cyclodextrin-bonded column. Journal of Chromatography B 802, 283–289. Fenn, J.B., Mann, M., Meng, C.K., Wong, S.F. and Whitehouse, C.M. (1989) Electrospray ionization for massspectrometry of large biomolecules. Science 246, 64–71. Harland, B.F., Smikle-Williams, S. and Oberleas, D. (2004) High performance liquid chromatography analysis of phytate (IP6 ) in selected foods. Journal of Food Composition and Analysis 17, 227–233. Henshall, A., Harrold, M.P. and Tso, J.M.Y. (1992) Separation of inositol phosphates by capillary electrophoresis. Journal of Chromatography 608, 413–419. Hsu, F.F. and Turk, J. (2000) Characterization of phosphotidylinositol, phosphatidylinositol-4-phosphate, and phosphotidylinositol-4,5-bisphosphate by electrospray ionization tandem mass spectrometry: a mechanistic study. Journal of the American Society for Mass Spectrometry 11, 986–999. Hsu, F.F., Goldman, H.D. and Sherman, W.R. (1990) Thermospray liquid-chromatographic mass-spectrometric studies with inositol phosphates. Biomedical and Environmental Mass Spectrometry 19, 597–600. Irving, G.C. and Cosgrove, D.J. (1981) The use of hypobromite oxidation to evaluate two current methods for the estimation of inositol polyphosphates in alkaline extracts of soils. Communications in Soil Science and Plant Analysis 12, 495–509. Irving, G.C. and Cosgrove, D.J. (1982) The use of gas–liquid chromatography to determine the proportions of inositol isomers present as pentakis- and hexakisphosphates in alkaline extracts of soils. Communications in Soil Science and Plant Analysis 13, 957–967. Kasim, A. and Edwards, H.M. (1998) The analysis for inositol phosphate forms in feed ingredients. Journal of the Science of Food and Agriculture 76, 1–9. Kerwin, J.L., Tuininga, A.R. and Ericsson, L.H. (1994) Identification of molecular-species of glycerophospholipids and sphingomyelin using electrospray ionization mass-spectrometry. Journal of Lipid Research 35, 1102–1114. L’Annunziata, M.F. and Fuller, W.H. (1971) Nuclear magnetic resonance spectra of acetate derivatives of soil and plant inositol phosphates. Soil Science Society of America Proceedings 35, 655–658. L’Annunziata, M.F. and Fuller, W.H. (1976) Evaluation of the mass spectral analysis of soil inositol, inositol phosphates, and related compounds. Soil Science Society of America Journal 40, 672–678.

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L’Annunziata, M.F., Gonzalez, J. and Olivares, L.A. (1977) Microbial epimerization of myo-inositol to chiro-inositol in soil. Soil Science Society of America Journal 41, 733–736. Llewelyn, J.M., Landing, W.M., Marshall, A.G. and Cooper, W.T. (2002) Electrospray ionization Fourier transform ion cyclotron resonance mass spectrometry of dissolved organic phosphorus species in a treatment wetland after selective isolation and concentration. Analytical Chemistry 74, 600–606. McKercher, R.B. and Anderson, G. (1968a) Characterization of inositol penta- and hexaphosphate fractions of a number of Canadian and Scottish soils. Journal of Soil Science 19, 302–310. McKercher, R.B. and Anderson, G. (1968b) Content of inositol penta- and hexaphosphates in some Canadian soils. Journal of Soil Science 19, 47–55. Neubauer, G. and Mann, M. (1999) Mapping of phosphorylation sites of gel-isolated proteins by nanoelectrospray tandem mass spectrometry: potentials and limitations. Analytical Chemistry 71, 235–242. Phillippy, B.Q. and Johnston, M.R. (1985) Determination of phytic acid in foods by ion chromatography with post-column derivatization. Journal of Food Science 50, 541–542. Reemtsma, T. and These, A. (2003) On-line coupling of size exclusion chromatography with electrospray ionization-tandem mass spectrometry for the analysis of aquatic fulvic and humic acids. Analytical Chemistry 75, 1500–1507. Sherman, W.R., Ackerman, K.E., Berger, R.A., Gish, B.G. and Zinbo, M. (1986) Analysis of inositol monophosphates and polyphosphates by gas-chromatography mass-spectrometry and fast-atom-bombardment. Biomedical and Environmental Mass Spectrometry 13, 333–341. Skoglund, E., Carlsson, N.-G. and Sandberg, A.-S. (1998) High-performance chromatographic separation of inositol phosphate isomers on strong anion exchange columns. Journal of Agricultural and Food Chemistry 46, 1877–1882. Steward, J.H. and Tate, M.E. (1971) Gel chromatography of soil organic phosphorus. Journal of Chromatography 60, 75–78. Taguchi, R., Hayakawa, J., Takeuchi, Y. and Ishida, M. (2000) Two-dimensional analysis of phospholipids by capillary liquid chromatography/electrospray ionization mass spectrometry. Journal of Mass Spectrometry 35, 953–966. Turner, B.L., Papházy, M.J., Haygarth, P.M. and McKelvie, I.D. (2002) Inositol phosphates in the environment. Philosophical Transactions of the Royal Society, London, Series B 357, 449–469.

4

Origins and Biochemical Transformations of Inositol Stereoisomers and Their Phosphorylated Derivatives in Soil Michael F. L’Annunziata The Montague Group, PO Box 5033, Oceanside, CA 92052-5033, USA

The determination of inositol phosphates in environmental samples is an analytical challenge that has hampered research on this topic for decades. The role of inositol phosphates in the environment, including the origins and biochemical transformations of the soil inositol phosphates and their detection by modern instrumental and isotopic techniques, was reviewed recently by Turner et al. (2002), and detailed reviews of the use of nuclear magnetic resonance (NMR) spectroscopy and mass spectrometry for the analysis of inositol phosphates in environmental samples also appear in this volume (see Murthy, Chapter 2, and Cooper et al., Chapter 3). This chapter summarizes early work (1967–1969) on the extraction and characterization of inositol phosphates in soil using a range of analytical procedures, including electron-impact mass spectrometry, infrared spectroscopy and proton NMR spectroscopy. It also describes subsequent work (1974–1977) conducted using isotopic-labelling techniques, and concludes with ideas on new approaches to research on the inositol stereoisomers and their phosphorylated derivatives in soils and other environmental samples using stable and radioactive isotopes.

Development of Methodologies for Characterization of Inositol Phosphates in Soil Extraction and preparative chromatography Inositol phosphates were extracted from leaf litter on the soil surface and the underlying soil organic matter of a forest and desert soil by the method of Anderson (1956). A flow chart of the extraction method is illustrated in Fig. 4.1. The precipitated fractions of extractable phosphoruslabelled precipitates A, B and C of Fig. 4.1 were combined, the phosphate groups were removed by hydrolysis in hydrochloric acid, the solution deionized with exchange resin and finally concentrated by evaporation to 25 ml. The phosphate groups were removed from the inositol rings to facilitate the separation and isolation of the inositol stereoisomers by preparative paper chromatography and their subsequent recrystallization and structural identification by a range of analytical techniques (L’Annunziata, 1970; L’Annunziata and Fuller, 1971a; L’Annunziata et al., 1972). The objective of the

©CAB International 2007. Inositol Phosphates: Linking Agriculture and the Environment (eds B.L. Turner, A.E. Richardson and E.J. Mullaney)

41

42

M.F. L’Annunziata

Soil Wash with 0.2 M HCI Extract with 1 M NaOH Centrifuge

Soil Wash with 1 M NaOH Centrifuge Supernatant

Supernatant 200 ml HAc Acidify with HCI to pH 0.5 Centrifuge

1. Precipitates A, B, C combined 2. Refluxed 2 days with 1 l of 1:1 conc. HCI 3. Deionized with exchange resin 4. Evaporated to 25 ml volume

Humic acid (discard) 5% of extractable P Raise pH to 8.5 with NH4OH Centrifuge

Supernatant

Precipitate A 75% of extractable P 2 l of 10% BaAc and centrifuge

Supernatant

Supernatant

Precipitate B (Ba phosphates)

Vacuum distillation to concentrate

Supernatant

Precipitate C (Ba phosphates)

Fig. 4.1. Flow chart illustrating the method of Anderson (1956) used to extract phosphorus from soil and plant samples. When used to extract phosphorus and its compounds from leaf litter, the word ‘soil’ in the flow chart should be replaced with ‘plant matter’.

study was to investigate the origins of the soil inositol phosphate stereoisomers by assessing the stereochemistry of inositols in leaf litter on the soil surface and in the soil organic matter of two soil types, a desert and a forest. The extracted phosphates were subjected to acid hydrolysis to remove all phosphate groups: a major portion of the hydrochloric acid was removed by vacuum distillation, anions and cations were removed by washing the extract through a column of deionizing resin and the aqueous solution concentrated to 25 ml by evaporation in an open beaker on a hot plate at moderate heat (60–80ºC). The inositols in the solution were separated by descending preparative paper chromatography by spreading ~0.4 ml of the solution onto individual large

(56 × 23 cm) chromatogram sheets under the warm air of a hairdryer. Approximately 50 chromatograms were required to isolate all of the inositol stereoisomers in the 25 ml concentrated extract. More details on the extraction and chromatographic procedures are available in L’Annunziata (1970) and L’Annunziata and Fuller (1971a). The inositol stereoisomers were eluted from the paper with hot deionized water and recrystallized in aqueous acetone or ethanol when possible (i.e. whenever sufficient sample was obtained). The inositol stereoisomers were identified by electron-impact mass spectrometry, infrared spectroscopy and solution 1H-NMR spectroscopy of the free inositols and their hexacetate derivatives.

Inositol Stereoisomers

Electron-impact mass spectrometry The electron-impact mass spectra of the inositol stereoisomers (myo- and D-chiro-inositol) were identical as expected, but the mass spectra provided initial evidence for the basic inositol structure by the molecular ion peak at m/e 180, the characteristic base peak at m/e 73 (C3H5O+2) and other characteristic ion peaks at m/e 102 (C4H6O3+ ) and m/e 60 (C2H4O2+ ). These are discussed in more detail below. The electron-impact mass spectrometers available at the time caused strong ionization and fragmentation and consequently a weak molecular ion peak. However, the fragmentation pattern together with the molecular ion peak was indispensable in making unequivocal identification of the compound as inositol or one of its derivatives. The mass spectral fragmentation of inositol, inosose and inositol-related compounds were studied in detail, the molecular formulas of the major ion peaks determined and the mass spectral ion fragmentation scheme postulated (L’Annunziata, 1970; L’Annunziata and Fuller, 1976). The mass difference of the major ion fragments was demonstrated to be a useful indicator of specific structural changes of the inositols. Typical electron-impact mass spectra obtained for the inositol stereoisomers (myo- and D-chiro-) and their natural products DL-epi-inosose and quebrachitol (monomethyl ether of L-chiroinositol) are illustrated in Fig. 4.2. A detailed study was made to identify the mass spectral ion fragments and the fragmentation scheme of the inositols by means of mass spectral peak shifts that resulted following deuterium labelling of the hydroxyl groups of the inositol, inosose and monomethyl ether (L’Annunziata, 1970; L’Annunziata and Fuller, 1976). The structures of the mass spectral ion fragments were postulated using the molecular formulas of the fragment ion peaks in conjunction with evidence from peak shifts produced by the mass spectra of molecular derivatives of inositol (e.g. inosose, quebrachitol and inositol hexacetate) and the deuteriumlabelled analogues of the derivatives. The molecular formulas were obtained by high-resolution mass spectrometry, or postulated for ion fragments of low mass by the elimination of unlikely combinations of carbon, hydrogen and oxygen listed in mass and abundance tables (Beynon and Williams, 1963; Buchs et al., 1968; L’Annunziata,

43

1970). The major mass spectra fragment ions and fragmentation pathway of myo-inositol is illustrated in Fig. 4.3 (L’Annunziata, 1970; L’Annunziata and Fuller, 1976). The mass spectra of the DL-epi-inosose helped to elucidate the mass spectral fragmentation scheme of myo-inositol, but its mass spectra also serve as a useful future reference for the identification of the inosose in soil systems, because the cyclic ketone stereoisomers have been previously proposed to occur as phosphorylated and non-phosphorylated intermediates in soil inositol and inositol phosphate epimerization reactions (e.g. L’Annunziata and Fuller, 1971a; Cosgrove, 1972; L’Annunziata, 1975). The ion fragment of m/e 163 observed in the mass spectrum of myo-inositol was of low intensity (0.1% of base peak); however, the difference in mass between this ion fragment and the protonated molecular ion of m/e 181 (Fig. 4.2a) was 18 mass units. This is equivalent to the loss of a water molecule, a common initial fragmentation pathway for polyhydroxy compounds. The ion fragment at m/e 144 is equivalent to the loss of 36 mass units or two water molecules. The mechanism for this elimination of water remains uncertain (Reed et al., 1962), although the three possible 1,2-, 1,3- and 1,4-eliminations may be considered. Two consecutive 1,2,-eliminations are illustrated in Fig. 4.3, resulting in the formation of ions II and III of m/e 163 and 144, respectively. A peak shift of two mass units from m/e 144 (C6H8O4+) to m/e 146 occurred in the mass spectrum of myo-inositol after deuterium labelling of the hydroxyl groups (C6H6(OD)6 of Fig. 4.2b), indicating the presence of two hydroxyl protons on the ion of m/e 144. This was explained by a keto–enol conversion between ions III and IV (Fig. 4.3) in which ion IV would shift to m/e 146 for the deuterium-labelled inositol. A major ion fragment at m/e 102 (C4H6O+3 ) contains two or three hydroxyl groups as indicated by its shift to m/e 104 and 105 after deuterium labelling (Fig. 4.2b). The formation of the fragment ion at m/e 102 (ion V of Fig. 4.3) may be expressed by the loss of a neutral ketene molecule from the ion of m/e 144 (ion IV of Fig. 4.3). The loss of ketene is characteristic of cyclic ketones. The keto–enol forms of ions V and VI of m/e 102 (Fig. 4.3) can explain peak shifts of two and three mass units of the deuterium-labelled hydroxyl groups.

44

M.F. L’Annunziata

OH

90

% of base peak

73 (C3H5O2+)

OH

OH

100

H

H OH

80 70

H

60

H OH

H

myo -Inositol,C6H6(OH)6

H

OH

50

m/e 163: 0.1% m/e 181: 0.1%

+ 60 (C2H4O2 )

40 30

+ 102 (C4H6O3 )

20 10

(a)

20

30

40

50

60

70

80

90

100

110

144 (C6H8O4+) 120

130

140

75 (C3H3D2O2+)

100

% of base peak

90 80

myo -Inositol-d6,C6H6 (OD)6

70 60 50 40 62 (C2H2D2O2+)

30 20

+ + 104 (C4H4D2O3 ) 105 (C4H3D3O3 )

10

(b)

20

30

40

HO

OH

80 70

70

80

90

100

110

120

130

140

150

=O 60

H

D,L-epi -Inosose,C6H5O (OH)5

OH

HO

60

60

73 OH

90

% of base peak

50

H

100

+ 146 (C6H6D2O4 )

H

H

H

40 30 20

OH C

H

H H .. + .O=C=C−C . OH H H .. O=C−C=C + OH

50

C HO

71

+ m/e 160: 2.4% (M + - H2O) m/e 179: 1.5% (MH+)

H C OH 102

+

142 (M− 2 H2O)

89

10

(c)

20

30

40

OH

100

% of base peak

90

H

80 70

H

60

H

50

70

80

100

130

140

m/e 158: 3.5 % m/e 194: 0.1% m/e 195: 0.2% 116 (C5H8O3+)

60 (C2H4O2+)

+ 102 (C4H6O3 )

H

OH

120

110

85 (C4H5O2+)

OH

OH

90

+ 73 (C3H5O2 ) 87 (C4H7O2+) Quebrachitol, C H (OH) OCH 6 6 5 3

H CH3O H

OH

60

50 40 30

+ 144(C6H6O4 )

20 10

(d)

20

30

40

50

60

70

80

90

100

110

120

130

140

150

m/e

Fig. 4.2. Electron-impact mass spectra of (a) myo-inositol; (b) myo-inositol-d6, C6H6(OD)6; (c) inosose; and (d) quebrachitol. (From L’Annunziata, 1970; L’Annunziata and Fuller, 1976.)

The base peak (most intense peak) at m/e 73 (C3H5O2+, Fig. 4.2a) in the mass spectra of inositol is produced by an ion fragment with two hydroxyl groups indicated by its shift to m/e 75 after deuterium labelling as illustrated in

DL-epi-

Fig. 4.2b. The ion is illustrated as the ring structure VII of Fig. 4.3 or as the resonance-stabilized allylic ion VII' in Fig. 4.4. This ion fragment (VII or VII') is also responsible for the base peak in the mass spectra of inosose and quebrachitol.

Inositol Stereoisomers

+

45

+

OH

+

OH

HO

OH

HO

OH OH

−H2O

OH

OH

−H2O

OH

HO

OH

OH OH

OH

I, m/e 181

II, m/e 163

III, m/e 144

+ HO

+ O

OH

+ O

OH

OH

−CH2 = C = O

OH OH

OH

VI, m/e 102

−C

V, m/e 102

IV, m/e 144

H

2

O −C H − H

O

=

C

=

O +

+

H

−CH•

H

HO

H

H C C

OH

HO

OH

VII, m/e 73 VIII, m/e 60 Fig. 4.3. The major electron-impact mass spectral fragmentation ions and fragmentation pathway of myo-inositol. (From L’Annunziata, 1970; L’Annunziata and Fuller, 1976.)

H C

H C

+ C

H

H

H C

H

H

+

H

C

HO

OH

HO

+ C

HO

(a)

(b)

VII′, m/e 73

H + O

OCH3

HO

IX, m/e 87

H

+

C OCH3

IX′, m/e 87

+ .. .. H O OCH3

X, m/e 85

C

H

H

OH

H C

H H C C C+

O

H H O C C C + ..

OCH3

X′, m/e 85

OCH3

HO

OCH3

OCH3

OH

OH

XI, m/e 116 XII, m/e 116 (c) (d) Fig. 4.4. Major ion fragments (a) base peak at m/e 73 (C3H5O2+) of the electron-impact mass spectrometry of myo-inositol, DL-epi-inosose and quebrachitol; (b) the second most intense peak at m/e 87 (C4H7O2+) at 98.9% of the base peak from the mass spectrum of quebrachitol; (c) the third most intense peak at m/e 85 (C4H5O2+) at 83% of the base peak from the mass spectrum of quebrachitol; and (d) an intense peak at m/e 116 (C5H8O3+) from the mass spectrum of quebrachitol. (From L’Annunziata, 1970; L’Annunziata and Fuller, 1976.)

46

M.F. L’Annunziata

The second-most predominate peak in the mass spectra of inositol and inosose and also a predominant peak in the mass spectra of quebrachitol is that at m/e 60 (C2H4O+2 ), which contains two hydroxyl groups (ion VIII of Fig. 4.3) as evidenced by its shift to m/e 62 following deuterium labelling of the hydroxyl groups. The mass spectra of quebrachitol labelled with deuterium in the hydroxyl groups [C6H6(OD5)OCH3] and of the hexacetate derivatives of myo-inositol [C6H6(OAc)6] and quebrachitol [C6H6(OAc5)OCH3] provided further evidence for the ion fragments and fragmentation scheme described (L’Annunziata, 1970; L’Annunziata and Fuller, 1976).

Infrared spectroscopy The infrared spectra provided perfect fingerprints for the myo- and chiro-inositol stereoisomers at every absorption frequency (e.g. for the ponderosa pine needles and forest soils; Fig. 4.5). The infrared spectra can serve as a tool in distinguishing specific inositol stereoisomers as evidenced by the dissimilarities between spectra of the myo- and chiro-inositol standards. These occur primarily in the ‘fingerprint’ region between 1350 and 800 cm⫺1. For example, marked dissimilarities are evident at 900 and 930 cm⫺1 for myo-inositol (left panel of Fig. 4.5) and at 870 and 905 cm⫺1 for chiro-inositol (right panel of Fig. 4.5). These may be due to differences in C–H rocking frequencies (Dyer, 1965). Also, marked dissimilarities are seen by four relatively sharp absorption frequencies at 1000, 1075, 1110 and 1150 cm⫺1 in the infrared spectrum of myo-inositol, which appear as two broad absorption frequencies at 1010 and 1100 cm⫺1 in the infrared spectrum of chiro-inositol. These are assigned to differences in C–O stretching frequencies (Lambert et al., 1998). The strong absorptions assigned to O–H stretching frequencies at 3300 cm⫺1 and the weaker absorption at 2900 cm⫺1 assigned to C–H stretching frequencies were common in all inositol infrared spectra studied. The very sharp truncated absorption peak at 1600 per cm seen in all the infrared spectra is that of a 0.07 mm polystyrene film used as a reference standard.

Proton nuclear magnetic resonance spectroscopy The 1H-NMR spectra provided strong evidence for the existence of inositol stereoisomers (Fig. 4.6). The most powerful spectrometers available to us at the time were 60 and 100 MHz instruments, so relatively large samples (>5 mg) were needed to obtain spectra in deuterium oxide with specially designed micro-NMR tubes. The myo-inositol standard and myo-inositols isolated from plant litter or underlying soil gave NMR spectra similar to those illustrated in the left panel of Fig. 4.6, with complex chemical shifts at 5.99, 6.31, 6.40, 6.49, 6.59, 6.70 and 6.78τ. Spectra of the chiro-inositol standard and the chiro-inositols isolated from plant litter and underlying soil differed from the myo-inositol spectrum by yielding complex chemical shifts at 5.97, 6.00, 6.31, 6.33, 6.38 and 6.42τ at different relative intensities than those for myo-inositol (right panel of Fig. 4.6). The 1H-NMR spectra of the hexacetate derivatives of the myo- and chiro-inositols isolated from the soil and plant litter were also perfect fingerprints of authentic myo- and chiro-inositol (L’Annunziata, 1970; L’Annunziata and Fuller, 1971b). Up to 88% yields were obtained with the acetylation of inositol samples as small as 20 mg. The increase in the molecular weight of the inositol upon acetylation was advantageous due to the very small amounts of inositols isolated from plant or soil. The NMR spectra of the acetate derivatives (not illustrated here) permitted the observation of the number of axial and equatorial acetoxy groups as sharp peaks due to the acetoxy protons (CH3CO2–) lacking the complexities produced by proton spin coupling.

Identification of D-chiro-inositol The optical rotation of the chiro-inositol enantiomer from the desert soil–plant system was determined to be +56º measured by a recording polarimeter (L’Annunziata, 1970). At about the same time, Cosgrove (1969) also reported the Dchiro-inositol as a hexakisphosphate. Further evidence of the D-chiro-inositol was subsequently provided by measurement of the positive optical

Wavelength 3

4

5

6

7

(microns)

8

9

10

Wavelength 11

12

13

14

0.0

15 0.0

0.30 0.40 0.50 0.60 0.70 1.0 1.5 ∞ 4000 3000

0.30 0.40 0.50 0.60 0.70 1.0 1.5 ∞

2000

1500

1200 cm

900

800

700

6

7

8

(microns) 9

10

11

12

13

14

−1

L-chiro-Inositol

0.10

0.10

0.20

0.20

0.30

0.30

0.40

0.40

0.50 0.60 0.70

0.50 0.60 0.70

1.0 1.5 ∞ 4000 3000

Inositol isomer from the forest soil

15 0.0

2000

1500

1200

1000

900

800

700

1.0 1.5 ∞

cm−1

3

4

5

6

7

Wavelength 8 9

(microns) 10 11

12

13

14

0.0

Inositol isomer from chromatographic spot b2

(b) 0.10

15 0.0

0.10

(c)

Inositol isomer from P. ponderosa needles

Absorbance

INFRACORD ⫻ 1371282

0.20

0.20

0.30

0.30

0.40 0.50 0.60 0.70

0.40 0.50 0.60 0.70

1.0 1.5 ∞ 4000 3000 INFRACORD ⫻ 1371282

2000

1500

1200

1000

900

800

700

Inositol Stereoisomers

(b)

1000

Absorbance

0.20

5

(a)

0.10

0.20

4

0.0

myo-Inositol

(a) 0.10 Absorbance

3

1.0 1.5 ∞

cm−1

Fig. 4.5. Characteristic infrared spectra of myo- and chiro-inositol. The left panel shows spectra of myo-inositol obtained from (a) a standard sample and (b) purified samples from a forest soil and (c) ponderosa pine needles from the soil surface. The right panel shows spectra of (a) a chiro-inositol standard and (b) the chiro-inositol isomer isolated from a desert soil or its velvet mesquite leaf litter on the soil surface. Regardless of source (i.e. plant or soil) the infrared spectra of pure myo-inositol or chiro-inositol were exact fingerprints. (From L’Annunziata, 1970; L’Annunziata and Fuller, 1971a; L’Annunziata et al., 1972.) 47

48

2.0 1000 500 250 100 50

400

H

6.49τ

200

300

100 H HO

H2O

H

OH OH

D

3.0

1000 500 250 100 50

Hz

H

4.0

400

HO H

7.0

300

9.0

200

H

H H 6.38t

DDS 6.70t 6.78t

5.97t

10

100

D

HO OH

6.00t

H CPS

H OH

H2O

myo - Inositol

5.99t 6.40

8.0

OH

H OH

HO

5.0 PPM(t) 6.0

OH H

H

OH

H

L-chiro-inositol 6.42t

6.31t

6.59t 6.31t

(a) 1000 500 250 100 50

400

300

200

6.49t

100

H D

Hz

8.0

7.0

2.0

3.0

1000 500 250 100 50

400

6.0

5.0 PPM(d) 4.0

4.0

5.0 PPM(t) 6.0 300

6.33t

200

3.0

2.0

7.0

8.0

1.0

0

9.0

10

100 D

H CPS

Forest soil inositol H2O

H2O 5.99τ

6.70τ

Inositol isomer from chromatographic spot b2

DDS TMS

7.0 6.0 5.0 PPM(δ) 4.0 3.0 2.0 1.0 0 (b) (b) 8.0 Fig. 4.6. Characteristic proton nuclear magnetic resonance (NMR) spectra of myo- and chiro-inositol. The left panel shows 100 MHz NMR spectra of myoinositol obtained from (a) a standard sample and (b) a purified sample from a forest soil. Peaks due to DDS internal standard are marked. The right panel shows 60 MHz NMR spectra of (a) a chiro-inositol standard and (b) a chiro-inositol isomer isolated from the soil or its leaf litter on the soil surface. Regardless of source (i.e. plant or soil) the NMR spectra of pure myo-inositol or chiro-inositol were exact fingerprints. (From L’Annunziata, 1970; L’Annunziata and Fuller, 1971a.)

M.F. L’Annunziata

(a)

Inositol Stereoisomers

rotation of the chiro-inositol isolated from the forest soil (L’Annunziata et al., 1972).

Identification of muco-inositol Another inositol stereoisomer, identified as mucoinositol, was isolated from velvet mesquite leaf litter on the soil surface, but was not identified in the desert soil. The basic inositol structure was confirmed by the characteristic infrared spectra of the inositol and of its hexacetate derivative. No standard NMR spectrum was available for the unequivocal identification of muco-inositol, but its major 1H-NMR absorption peak at 1.53 ppm was in very close agreement to the 1.52 ppm for muco-inositol reported by Brownstein (1959). The melting point (m.p.) of the hexacetate derivative (175.5–177ºC) was in close agreement with that of authentic muco-inositol hexacetate (177–178ºC) reported by Nakajima et al. (1959). muco-Inositol occurs in higher plants (Richter et al., 1990; Wanek and Richter, 1995; Peterbauer and Richter, 1998), but neither muco-inositol nor its phosphorylated form has been reported in soil. Further discussion of the inositol phosphate stereoisomers can be found elsewhere (Turner, Chapter 12, this volume).

Significance of Inositol Stereoisomers in Soil Not all of the inositol stereoisomers present in the soil or plant samples were identified, due to the small amounts isolated and the limited power of the NMR spectrometers during the 1960s. scylloInositol and neo-inositol phosphates occur in soils (Turner et al., 2002) and may have been present in our samples, as indicated by a small amount of an unidentified compound in one of the paper chromatogram bands (see c and d of Fig. 6 in L’Annunziata, 1970, p. 38). However, it was not possible to recrystallize and run NMR spectra on that small sample. All chromatographic separations and mass spectrometry and NMR spectroscopic measurements were performed on acid hydrolysates of the phosphate fractions A, B and C of plant or soil extracts according to the method of Anderson (1956) illustrated in Fig. 4.1. Hydrolysis

49

of the phosphates was performed by refluxing the precipitates A, B and C in 1:1 hydrochloric acid for 2 days, followed by deionization with exchange resin to remove inorganic ions to yield aqueous solutions of the free inositols. At the time of these studies (1967–1969) only electron-impact mass spectrometry was available, which required samples in crystalline form that could be volatilized with heating in the mass spectrometer followed by electron impact with molecules in the vapour phase. Consequently, mass spectrometry of inositols in the phosphorylated form was not possible, so the phosphate groups were removed to facilitate identification of the inositol stereoisomers. Likewise, only 60 or 100 MHz NMR spectrometers were available, which required the dissolution of the pure crystalline inositol in deuterium oxide. The stability of the inositols under acid hydrolysis was tested by analysing myo-inositol by 1H-NMR spectroscopy before and after treatment with 20% deuterated hydrochloric acid under reflux, but no observable change occurred after 3 days. The studies described above were intended to investigate only the organic carbon stereochemistry (inositol moiety) of the phosphates by mass spectrometry and NMR spectrometry. Subsequently, Cosgrove (1980) reported that considerable amounts of polysaccharide and nitrogenous material could exist in the precipitate fractions isolated by the method of Anderson (1956), raising the possibility that some unphosphorylated inositols may be extracted along with the phosphorylated forms, particularly in extracts of plant matter. Ion-exchange chromatographic separation of inositol phosphate fractions prior to acid hydrolysis would therefore be required to conclude irrefutably that the inositols are isolated only in the phosphorylated form. The fact that both the myo-inositol and D-chiro-inositol were identified in the plant litter on the soil surface and in the underlying soil pointed to plant residue as a possible origin of the soil inositol phosphate stereoisomers. However, soil microbial epimerization of myo-inositol to D-chiro-inositol was suggested as another possible origin of this stereoisomer (L’Annunziata, 1970) in light of the only structural difference between the two stereoisomers, which is the stereochemistry at a single carbon atom (L’Annunziata, 1970; Fig. 4.7). The epimerization of myo-inositol to D-chiro-inositol involving a cyclic ketone

50

M.F. L’Annunziata

1

2

1 3

6

myo-Inositol (1,2,3,5/4,6)-Inositol

− 2 O

3

6

4 5

H

2

1 6

3

4

5

5

4 (D)-chiro-Inositol (1,2,4/3,5,6)-Inositol

Inosose intermediate

Fig. 4.7. The proposed microbial epimerization of myo-inositol to D-chiro-inositol in soil. (From L’Annunziata, 1970.) Numbering follows International Union of Pure and Applied Chemistry (IUPAC) nomenclature.

intermediate in Trifolium incarnatum was observed by Scholda et al. (1964), but there was no evidence then of any such reaction occurring in soils. Subsequent work was therefore undertaken with the radioisotope 14C to provide unequivocal evidence for the soil microbial epimerization of myo-inositol to D-chiro-inositol.

Isotopic Techniques for Studying Inositol Phosphates in Soil When myo-inositol uniformly labelled with 14C [14C(U)-myo-inositol] became available, L’Annunziata et al. (1975) proposed that it could be used to elucidate the pathways of myo-inositol transformations in soil. The only stereoisomers of inositol known to occur in soils as phosphorylated forms were the myo-, D-chiro-, neo- and scylloinositol. The latter three differ from myo-inositol (the most abundant of the isomers in soil, plant and animal systems in phosphorylated or free form) only at one carbon atom (Fig. 4.8). This suggested the possibility that D-chiro-, neo- and scyllo-inositol or their phosphates originated in soils by the epimerization of myo-inositol or its phosphate at one of the carbon atoms marked by symbols in Fig. 4.8. To find the fate of myo-inositol and its hexakisphosphate in soils, studies were initiated with 14C isotope tracer. A study of the soil microbial metabolism of uniformly labelled 14C(U)-myo-inositol, 14 C(U)-myo-inositol hexakisphosphate and 14C(U)iron(III) phytate was carried out. The labelled compounds were incubated in either microbially active or sterilized forest soil in Bartha and Pramer (1965) incubation flasks (Fig. 4.9). The soil was taken from the A1 horizon of a red forest soil

(Andisol), had a pH of 5.8 and contained 10.9% organic matter and 18% clay (L’Annunziata and Gonzalez, 1977). The flasks permitted a determination of the soil microbial metabolism rates by measurement of the evolution rates of 14CO2 trapped by 0.1 M KOH in an attached side arm. The results indicated that 61% of the carbon in myo-inositol, 1.9% of the carbon in myo-inositol hexakisphosphate and none of the carbon in iron(III) phytate were oxidized to carbon dioxide on 12 days of incubation (Table 4.1; L’Annunziata and Gonzalez, 1977). No carbon was oxidized to

1

2

2 3

6

1 6

3

4

5

5

4

myo -Inositol (1,2,3,5/4,6)-Inositol

D-chiro-Inositol (1,2,4/3,5,6)-Inositol

2 1

1

2

6

3 5

3

6

4

4 5

neo -Inositol (1,2,3/4,5,6)-Inositol

scyllo-Inositol (1,3,5/2,4,6)-Inositol

Fig. 4.8. The soil inositol stereoisomers. The stereochemical differences of the D-chiro-, neo- and scyllo-inositol stereoisomers or their phosphates in comparison with the myo-inositol stereoisomer are indicated by the symbols ●, ■ and ▲, respectively. (Adapted from L’Annunziata, 1975.)

Inositol Stereoisomers

e

d

f h g

b c a

i

Fig. 4.9. A Bartha and Pramer incubation flask. The components and contents of the flask are as follows: (a) 50 g soil moistened to 70% field capacity or other suitable moisture content; (b) 250 ml Erlenmeyer flask fused to (c) a 50 ml test tube with round bottom; (d) Ascarite (NaOH-coated silica) absorber of atmospheric carbon dioxide; (e) rubber stopper; (f) stopcock to permit incoming air when opened and (e) rubber stopper removed during sampling of KOH; (g) 15 cm long 15-G needle with short length of polyethylene tubing extending to the bottom of the round base of the side arm; (h) rubber policeman cap, which is replaced with a calibrated syringe to allow for the injection of (i) 10 ml of 0.1 M KOH solution into the side arm and to allow for the removal of the KOH solution and exchange with fresh KOH solution for the measurement of 14CO2 at periodic intervals. (From Bartha and Pramer, 1965. Reprinted with permission from Lippincott Williams & Wilkins.)

The rate of oxidation of 14C-labelled myo-inositol hexakisphosphate to carbon dioxide dropped rapidly from 0.99% after day 1 to 0.17% after day 2, and then to 0.05% after day 12 (Table 4.1), indicating a rapid fixation of this compound

51

into a form unavailable to microorganisms (L’Annunziata and Gonzalez, 1977). The soil incubated with 14C-myo-inositol under normal and sterile conditions was fractionated according to Fig. 4.1. The quantities of 14C found in each soil fraction for both the non-sterile and sterile soils are shown in Table 4.2 (L’Annunziata and Gonzalez, 1977), which lists the percentage of the total radiocarbon remaining in the soil after 60% and 0% of the radioisotope label had evolved as 14CO2 for the non-sterile and sterile treatments, respectively. The quantity of 14C encountered in each fraction is greatly dependent upon non-sterile or sterile soil conditions. As indicated in Table 4.2, fractionation of the soil incubated with 14C(U)-myo-inositol under nonsterile conditions yielded small quantities of radioisotope in the water- and HCl-extractable fractions. Larger quantities of 14C were encountered in the unextractable and the humic-acid fraction. The unextractable fraction may consist of clay-bound inositol and its metabolites. The phosphate fractions E, F and G were previously shown to contain myo-inositol hexakisphosphate (Anderson, 1956; L’Annunziata, 1970; L’Annunziata and Fuller, 1971a). Phosphate fractions E, F and G of Table 4.2 (notation used in the writer’s paper of 1977) are the same as fractions A, B and C of Fig. 4.1. Significant quantities of 14C were found in each of these fractions. Summing the percentages of the 14C in each of the fractions with the value of 61.0% for the radiolabel evolved as 14 CO2 (Table 4.1) yielded a total of 100.72%. This accounts quantitatively for all of the 14C(U)-myoinositol applied to the non-sterile soil. The phosphate fractions E, F and G were submitted to hydrochloric acid hydrolysis to remove the phosphate groups, and 14C-myo-inositol was identified in the hydrolysed-phosphate fractions. The soil-bound inositol carbon predominates under non-sterile conditions (Table 4.2). The adsorption of inositol by clay minerals has been demonstrated (Greenland, 1956). The high mineral adsorption of 14C (31.5% soil-bound or unextractable) in the non-sterile soil is considered to be caused by the adsorption of microbial metabolites. The predominance of metabolites of 14 C-myo-inositol in the non-sterile soil was estimated on the basis of the relatively high evolution of 14CO2 under non-sterile conditions. The humic acid-bound inositol carbon formed under

52

M.F. L’Annunziata

Table 4.1. Daily evolution of 14C carbon dioxide from an Andisol sandy loam treated with uniformly labelled 14C myo-inositol, 14C phytic acid or 14C iron(III)-phytate. (From L’Annunziata and Gonzalez, 1977.) 14

14

C-myo-inositol (non-sterile)

14

Day 1 2 3 4 5 6 7 8 9 10 11 12 Total

C-CO2 (%)a

38.25 4.49 3.63 2.80 2.17 1.77 1.67 1.32 1.44 1.45 0.98 0.99 61.0 ± 0.44%b

C-myo-inositol (sterile)

Total CO2 (meq)

14

14

C-CO2 (%)a

Total CO2 (meq)

− − − − − − − − − − − − −

− − − − − − − − − − − − −

0.644 0.493 0.486 0.460 0.414 0.424 0.367 0.384 0.424 0.454 0.325 0.504 5.38 ± 0.42%b

14

C-phytic acid

14

C-CO2 (%)a

0.99 0.17 0.14 0.12 0.09 0.07 0.07 0.05 0.05 0.06 0.04 0.05 1.90 ± 1.4%b

C-iron(III) phytate

Total CO2 (meq)

14

C-CO2 Total CO2 (%)a (meq) − − − − − − − − − − − − −

0.716 0.480 0.489 0.472 0.453 0.375 0.379 0.292 0.318 0.382 0.286 0.350 4.99 ± 0.14%b

0.598 0.440 0.429 0.447 0.397 0.370 0.374 0.317 0.316 0.368 0.268 0.314 4.64 ± 2.6%b

a

Percentage of radiolabel applied to the soil. Variation between duplicate measurements or the average deviation expressed as a percentage of the mean.

b

non-sterile soil conditions was abundant, although only a trace was encountered in the sterile soil (Table 4.2). The low level of 14C in the humic acid fraction of the sterile soil indicates the importance of microbial activity on interactions between humic acid and inositol and its metabolites.

The water-extractable fraction of the soil incubated with 14C(U)-myo-inositol was submitted to descending paper chromatography on large (56 × 23 cm) chromatogram sheets, as described previously, and the paper chromatogram subjected to autoradiography to visualize the location of the separated 14C-labelled compounds

Table 4.2. Phosphorus and radiocarbon contents in various fractions of soil incubated with uniformly labelled 14C-myo-inositol under sterile and non-sterile conditions. (From L’Annunziata and Gonzalez, 1977.) Phosphorusa (ppm) Fraction Soil-bound H2O-extractable HCl-extractable Humic acid Phosphates E Phosphates F Phosphates G Residue Total a

14

C-labelb (%)

Organic

Inorganic

Non-sterile

Sterile

− − − −c 0.50 × 104 3.69 × 103 8.12 × 103 −c 1.68 × 104 ± 3.1%d

− − Trace 8.32 × 105 1.04 × 104 4.56 × 103 1.08 × 103 −c 8.48 × 105 ± 4.1%d

31.54 1.36 4.54 22.00 5.54 1.72 0.54 36.92 104.16 ± 2.5%d

0.42 85.83 13.90 0.05 0.15 0.03 0.12 1.01 101.51 ± 2.6%d

Analysis of the non-sterile soil. Percentage of total radiolabel remaining in the soil. c Undetermined because of high salt concentrations or interfering ions. d Variations between duplicate measurements or the average deviation expressed as a percentage of the mean. b

Inositol Stereoisomers

(L’Annunziata et al., 1977). A typical autoradiogram is illustrated in Fig. 4.10. Bands a1 and a2 of the water-extractable 14C of the sterile and non-sterile soil were demonstrated to be due to the original unmetabolized 14 C-myo-inositol. It is of much interest to note that the water-extractable fraction of the sterile soil contained only the original unmodified 14C-myoinositol. The second band (b1) in Fig. 4.10 was identified to be 14C-chiro-inositol by infrared spectrometry and thin-layer co-chromatography with standards (Fig. 4.11). The optical activity of the chiro-inositol was not measured, but it was assumed to be the D-chiro-enantiomer as L-chiroinositol has not been identified in soils. On the

O

a1

53

basis of the soil incubation studies with 14C(U)myo-inositol, the carbon and phosphorus pathways involving myo-inositol and its phosphates were proposed (Fig. 4.12).

Recommendations for Future Research with Stable and Radioactive Isotopes Studies with inositol and its phosphates labelled with radioactive isotopes, including 3H, 14C, 33P and 32P as single or dual isotope labels, offer the advantages of fast and easy real-time detection and

O

a2

b1

c1

d1

(a)

(b)

Fig. 4.10. Contact photographic prints made from X-ray film autoradiographs of paper chromatograms of (a) the water-extractable fraction of the non-sterile soil incubated with 14C(U)-myo-inositol and (b) the water-extractable fraction of the sterile soil incubated with 14C(U)-myo-inositol. (From L’Annunziata et al., 1977.) The chromatogram of (a) the non-sterile soil extract exhibited four bands marked a1, b1, c1 and d1, while the chromatogram of (b) the sterile soil extract exhibited only one band, marked a2. Bands a1 and a2 were demonstrated to be the original 14C-myo-inositol, while band b1 was demonstrated to be 14 C-chiro-inositol, a microbial metabolite of 14C-myo-inositol. Bands c1 (weak) and d1 were not identified.

54

M.F. L’Annunziata

OH OH HO OH HO

OH

[14C]-myo-inositol

cpm

OH

OH

HO OH HO OH

20

30

[14C]-chiro-inositol

SF 10

O

Fig. 4.11. A thin-layer chromatogram (TLC) of the eluted paper chromatographic 14C-bands a1 and b1 of the non-sterile soil extract with authentic myo- and chiro-inositols. (From L’Annunziata et al., 1977.) A recording of the 14C radioactivity (500 counts per minute full scale) originating from components applied to the origin on the TLC plate is positioned directly above the plate. The letters O and SF mark the origin and solvent front, respectively. The radioactivity of the chiro-inositol metabolite represented ~4% of that of its myoinositol precursor.

measurement of the isotope label. Flow-scintillation analysis of liquid chromatography effluents in homogeneous or heterogeneous flow cells provides efficient analysis of beta-particle-emitting radionuclide tracers (L’Annunziata, 2003). Highperformance liquid chromatography (HPLC) would be suitable mostly for the separation of the free non-phosphorylated inositol stereoisomers and their related compounds (e.g. non-phosphorylated inosose), such as those discussed in this chapter, which were subjected to acid hydrolysis to remove the phosphate groups prior to their chromatographic separation and structural elucidation.

Other separation techniques such as ion-exchange chromatography are currently the most efficient in separating the inositol stereoisomers in phosphorylated forms (see also Cooper et al., Chapter 3, this volume). The mobile phase in ion-exchange chromatographic separation systems may contain strong chemical components that could preclude the use of on-line mass spectrometry or NMR spectrometry. In such cases fraction collection of the separated components as determined by the flow scintillation detector would be required prior to the preparation of samples for mass spectrometry and NMR spectroscopy. Whenever on-line spectroscopic techniques are permitted the effluent from the flow scintillation analyser can be connected with an ultraviolet absorbance detector, mass spectrometer or NMR spectrometer for molecular structure analysis using chromatograph effluent splitting (Fig. 4.13). Such methods, often referred to as HPLC–FSA–UV–NMR–MS analysis systems, are described in detail elsewhere (L’Annunziata, 2003). The free inositols would not yield any ultraviolet absorption, so the ultraviolet detector would be superfluous in such cases; however, the expected inositol inosose (ketone) intermediates in the chromatograph effluent would likely produce ultraviolet absorption. Radiolabelled compounds in the effluent would be detected by flow scintillation prior to on-line mass spectrometry, NMR spectroscopy or fraction collection for offline spectroscopy. 1H, 31P or natural-abundance 13 C NMR may be used for molecular structure elucidation. The NMR spectra on-line are obtained using a stop-flow method with resonance signal acquisitions varying from several minutes to hours (Hansen et al., 1999; Smith et al., 1999; Bailey et al., 2000; Sweeney et al., 2000). The techniques are reviewed by L’Annunziata (2003). Both homogeneous and heterogeneous flow scintillation analyses of the column chromatograph effluents are possible. In homogeneous flow scintillation analysis a liquid scintillator is used to detect and measure the amount of radioactivity in the chromatograph effluent. In such a case the flow scintillation analyser is equipped with a splitter to permit a fraction of the effluent to go on to the mass spectrometer, NMR spectrometer or fraction collector. If ionexchange chromatographic separations of inositol phosphates do not permit on-line mass spectrometry or NMR due to strong chemical mobile

Inositol Stereoisomers

OH

H

H 2

(b) Clay−inositol carbon absorption

O

H

OH

(c)

H HO

Clay minerals

55

H

1

Humic acid−inositol carbon complexes or bonding

Humic acid OH

HO H

Acetobacter suboxidans Berman and Magasanik (1966a) and Posternak(1962)

H

myo -Inositol (1,2,3,5/4,6)-Inositol

O

(d)

H H O

(a) Epimerization

Soil phosphate

H

HO

HO

OH

H

OH

Inositol phosphate isomers

+3

Fe AI

+3

H

H

OH

O

H

H

OH

HO 2

H H

Dehydratase Berman and Magasanik (1966a)

(d)

H

H 1

(e) Soil phosphate

OH

H

(e)

O

OH OH

H

H H

(D)-chiro-Inositol (1,2,4/3,5,6)-inositol

O

H

H OH

O H

H2O Ring cleavage Iron and aluminium phytate (in acid soil)

Hydralase Berman and Magasanik (1966b) Other catabolic reactions

CO2

Fig. 4.12. Proposed carbon and phosphorus pathways of soil inositol including (a) the microbial epimerization of myo-inositol to (D)-chiro-inositol in soil. (From L’Annunziata and Gonzalez 1977.) The epimerization of myo-inositol to chiro-inositol was demonstrated by the use of 14C-labelling; however, the optical rotation of the chiro-inositol was not measured, hence the parenthesis around the dextrorotatory notation. Other pathways of myo-inositol carbon in soils illustrated include (b) clay mineral absorption; (c) humic acid formation; (d) oxidation to carbon dioxide; and (e) phosphorus fixation through phosphorylation and formation of iron and aluminium phytate complexes in acid soil.

phases, the effluent splitter can be used to direct a portion of the effluent to a fraction collector. The other fraction of the split effluent is mixed in liquid scintillator, radioactivity peaks plotted and count rates (counts per minute) converted to disintegration rates (disintegrations per minute) with prior radionuclide detection efficiency measurements. Reported detection efficiencies for radionuclide tracers in column chromatograph effluents by the homogeneous flow scintillation method are 20–60% for 3H, 70–95% for 14C, 70–95% for 33P and 85–95% for 32P. The heterogeneous flow scintillation method does not require a liquid scintillator because it uses a solid scintillator (e.g. SolarScint®, Trademark of PerkinElmer Life and Analytical Sciences) packed into fine Teflon tubing through which the chromatograph effluent flows. This means that heterogeneous flow scintillation analysis is less

cumbersome, less expensive and no chromatograph effluent splitting is required for radioactivity measurement. Again, a fraction collector could be used whenever off-line mass spectrometry and NMR spectrometry are desirable. Heterogeneous flow scintillation detection efficiencies are somewhat lower than the homogeneous method, being 3% for 3H and 30% for 14C. Much higher detection efficiencies would be expected for 32P, as this radionuclide emits a beta particle with maximum energy (Emax = 1710 keV), which is more than tenfold greater than that of 14C (Emax = 155 keV). Following are some of the questions that may be answered with radionuclide tracers: 1. Does the soil microbial epimerization of myoinositol also yield scyllo-inositol and neo-inositol; if so, does the epimerization occur via a mechanism similar to the epimerization of myo-inositol to chiro-inositol?

56

M.F. L’Annunziata

Chromatography and NMR spectrometer console NMR magnet

HPLC pump FSA detector and console

Injector

Column

254 UV detector

Splitter

Mass spectrometer console

HPLC−NMR probe

Mass spectrometer

Fig. 4.13. Instrumental set-up of the column chromatograph–ultraviolet–flow scintillation analysis–nuclear magnetic resonance–mass spectrometry apparatus. Where ion-exchange liquid chromatographic separations of inositol phosphates contain strong chemical mobile phases that do not permit on-line mass spectrometry or nuclear magnetic resonance (NMR) spectroscopy, the effluent splitter can be used to direct a portion of the effluent on to a fraction collector. (Modified from Hansen et al., 1999.)

2. What are the intermediates of the soil microbial epimerization of myo-inositol to chiro-inositol? 3. Do soil myo-inositol phosphates also undergo epimerization? 4. How stable are the alkaline earth (barium, calcium) salts or iron and aluminium salts of soil inositol phosphate stereoisomers as a function of soil pH and other factors, as the inositol phosphate salts, under certain soil conditions, may comprise a large pool of soil organic phosphorus not readily available to plants or soil microorganisms? 5. What are the rates of phosphorylation and dephosphorylation of inositol using 14C/32P dualisotope labels? The first three questions can be investigated with the 14C isotope followed by mass spectrometry and 1H, 13C or 31P NMR. The flow scintillation analyser, which may be coupled to an ionexchange chromatograph column, is equipped to discriminate and measure dual-isotope labels such as 3H–14C, 14C–32P and 33P–32P. The meas-

urement of single and dual radioisotope-labelled compounds in column chromatograph effluents is reviewed by L’Annunziata (2003). Examples of the possible soil microbial epimerization pathways (some yet unknown) of myo-inositol that may be resolved with the use of 3H or 14C isotope tracer techniques are illustrated in Fig. 4.14. Dual-labelled precursors such as uniformly ring-labelled 3H–14C(U)-myo-inositol and/or its phosphates together with the measurement of the 3H/14C isotope ratios of the myoinositol precursor and its product compound(s) can provide information on whether one or more atoms of 3H are lost in the process. This would provide insight into the mechanisms of myo-inositol transformations in soil. The following hypothetical case may be taken as an example. If uniformly ring-labelled 3H–14C(U)-myo-inositol of known 3H and 14C activities, measured in disintegrations per minute (DPM), is incubated in soil, and the 3H and 14 C activities of the product compound determined, we can deduce whether one or more atoms of 3H are lost in the biochemical transformation.

Inositol Stereoisomers

*2

*1 6

57

3

*

* 4

*

*

5

? ?

*

1

ial rob Mic

myo-Inositol (1,2,3,5/4,6)-Inositol

6

3

*

ep

3

*

4

*1

?

*2

6*

*1 *

5

*

*

oil

?

*

4

5*

*

*

O

6

*

(+)−Viboinosose intermediate st s

5

3

O

ore in f

*

*

2

tion

6

*

*1

*

4

5

riza

*2

*1

*

ime

O

*2

4

3

*

neo-Inositol (1,2,3/4,5,6)-Inositol

6

*

2

2

* *3

* 5*

4

3

*

* *4

*

1*

5

6

*

(D)-chiro-Inositol (1,2,4/3,5,6)-Inositol

scyllo-Inositol (1,3,5/2,4,6)-Inositol

Phosphorylation

*=

14C

or

3H

Inositol phosphate stereoisomers

Fig. 4.14. The soil microbial epimerization of myo-inositol to (D)-chiro-inositol and proposed possible epimerization pathways (marked with a question mark) of myo-inositol to the soil neo- and scylloinositols and their phosphates. The epimerization of myo-inositol to chiro-inositol was demonstrated by the use of 14C-labelling. (From L’Annunziata et al., 1977.) The optical rotation of the chiro-inositol was not measured in this experiment with 14C, although the dextrorotatory character of the chiro-inositol was previously reported. (From Cosgrove, 1969; L’Annunziata, 1970; L’Annunziata et al., 1972.) The (+)viboinosose intermediate illustrated above was identified to occur in Trifolium incarnatum by Scholda et al. (1964). The inosose has not been demonstrated to occur in soil systems. The asterisks can represent either 3H or 14C isotope labels.

Table 4.3. Example of a hypothetical application of the 3H to microbial transformation of myo-inositol. DPMa

Normalized DPM

14

C activity ratio measurements to the soil

Normalized 3H/14C ratio

3

H/14C activity ratio of uniformly ring-labelled 3H-14C(U)-myo-inositol precursor 3 H = 1200 (1200)(2.92) = 3500 14 C = 3500 (3500)(1) = 3500 3 H/14C activity ratio of metabolite (e.g. D-chiro-inositol) 3 H = 230 (230)(2.92) = 672 14 C = 800 (800)(1) = 800 a

DPM = disintegrations per minute.

1:1 or 6:6

672/800 or 5:6

58

M.F. L’Annunziata

OH OH

*

OH H

*

H

*

*

H

*

OH

H

H

*

H

*

OH

*

H 2

OH H

OH

(+)-Viboinosose (loss of one 3H atom)

:* = 6:6

H

*

*

H

myo-Inositol (1,2,3,5/4,6)-Inositol H

OH

o

*

H

OH

OH

or

*

2 H

OH

3H:14C

OH

OH

*

H 1

H

*

OH

H

*

*

OH

1 H

*

*

OH

H

OH

D-chiro -Inositol (1,2,4/3,5,6)-Inostiol 3H:14C = 5:6

Fig. 4.15. Epimerization of myo-inositol to D-chiro-inositol via (+)-viboinosose intermediate that involves the loss of a hydrogen atom from the inositol ring found in Trifolium incarnatum by Scholda et al. (1964). The encircled ring hydrogen atoms and asterisks represent the molecular locations of 3H and 14C isotope labels.

Table 4.3 provides hypothetical experimental data to illustrate the application. The 3H and 14 C activities of the uniformly ring-labelled myoinositol precursor are determined to be 1200 and 3500 DPM, respectively. Because there are six hydrogen and six carbon atoms on the inositol ring, it is necessary to first normalize the 3H/14C activity ratio of the myo-inositol precursor so that the ratio equals 1:1 or 6:6. This is done, in this example, by multiplying the 3H activity by the factor 2.92 necessary to bring up the 3H activity to equal that of the 14C and provide the required ratio as illustrated in Table 4.3. If the metabolite, e.g. D-chiro-inositol, has 3H and 14C activities of 230 and 800 DPM, respectively, we must first multiply the 3H activity by the same factor 2.92 used to normalize the precursor activity ratio. As illustrated in Table 4.3, this yields a 3H/14C ratio of 672 DPM/800 DPM or 5:6. From the change in the 3H/14C activity ratios from 6:6 to 5:6 we may conclude the loss of one hydrogen atom in the process and deduce an inosose intermediate such as that illustrated in Fig. 4.15. Activity ratio studies with dual-labelled 14C/32P-myo-inositol

phosphates may also be carried out, the possibilities of which are limited only by the imagination of the researcher. A review of the application of dual-radioisotope activity ratios in numerous studies of biochemical transformations is given by L’Annunziata (1984). Studies with 3H, 14C, 33P or 32P as single- or dual-labelled myo-inositol or its phosphates offer advantages of quick and facile tracing by flow scintillation analysis of soil microbial isotopelabelled metabolites in chromatograph effluents. The potential for the applications of isotopes in the study of inositol phosphate transformations is indeed significant. The work described here provides evidence for two origins of the inositol stereoisomers and their phosphates in soils, namely plant residues and epimerization of myo-inositol by soil microbes. Future studies with stable and radioactive isotopes can provide more information on the origins and transformations of the inositol stereoisomers and their role in the chemistry of soil phosphorus, soil fertility and soil environment.

References Anderson, G. (1956) The identification and estimation of soil inositol phosphates. Journal of the Science of Food and Agriculture 7, 437–444. Bailey, N.J.C., Cooper, P., Hadfield, S.T., Lenz, E.M., Lindon, J.C., Nicholson, J.K., Stanley, P.D., Wilson, I.D., Wright, B. and Taylor, S.D. (2000) Application of directly coupled HPLC–NMR–MS/MS to the identification of metabolites of 5-trifluoromethylpyridone (2-hydroxy-5-trifluoromethylpyridine) in hydroponically grown plants. Journal of Agriculture and Food Chemistry 48, 42–46. Bartha, R. and Pramer, D. (1965) Features of a flask and method for measuring the persistence and biological effects of pesticides in soils. Soil Science 100, 68–70. Berman, T. and Magasanik, B. (1966a) The pathway of myo-inositol degradation in Aerobacter aerogenes. Dehydrogenation and dehydration. Journal of Biological Chemistry 241, 800–806.

Inositol Stereoisomers

59

Berman, T. and Magasanik, B. (1966b) The pathway of myo-inositol degradation in Aerobacter aerogenes. Ring scission. Journal of Biological Chemistry 241, 807–813. Beynon, J.H. and Williams, A.E. (1963) Mass and Abundance Tables for use in Mass Spectrometry. Elsevier, New York. Brownstein, S. (1959) Shifts in NMR absorption due to steric effects. II. Polysubstituted cyclohexanes. Journal of the American Chemical Society 81, 1606–1608. Buchs, A., Charollais, E. and Posternak, T. (1968) Recherches dans la serie des cyclitols. XXXVII. Etude par spectrometrie de masse. Helvetica Chimica Acta 51, 695–707. Cosgrove, D.J. (1969) The chemical nature of soil organic phosphorus. II. Characterization of the supposed DLchiro-inositol hexaphosphate component of soil phytate as D-chiro-inositol hexaphosphate. Soil Biology and Biochemistry 1, 325–327. Cosgrove, D.J. (1972) The origin of inositol polyphosphates in soil. Some model experiments in aqueous systems involving the chemical phosphorylation of myo-inositol and the epimerization of myo-inositol pentaphosphates. Soil Biology and Biochemistry 4, 387–396. Cosgrove, D.J. (1980) Inositol Phosphates: Their Chemistry, Biochemistry, and Physiology. Elsevier, Amsterdam, The Netherlands. Dyer, J.R. (1965) Applications of Absorption Spectroscopy of Organic Compounds. Prentice-Hall, Englewood Cliffs, New Jersey. Greenland, D.J. (1956) The absorption of sugars by montmorillonite. I. X-ray studies. Journal of Soil Science 7, 319–328. Hansen, S.H., Jensen, A.G., Cornett, C., Bjørnsdottir, I., Taylor, S., Wright, B. and Wilson, I.D. (1999) High-performance liquid chromatography on-line coupled to high-field NMR and mass spectrometry for structure elucidation on constituents of Hypericum perforatum L. Analytical Chemistry 71, 5235–5241. L’Annunziata, M.F. (1970) Soil–plant relationships and spectroscopic properties of inositol stereoisomers; the identification of D-chiro- and muco-inositol in a desert soil–plant system. PhD thesis, University of Arizona, Tucson, Arizona. Available from University Microfilms International (UMI)/Bell and Howell Information and Learning (Dissertation Order No. 7104237), Ann Arbor, Michigan. L’Annunziata, M.F. (1975) The origin and transformations of the soil inositol phosphate isomers. Soil Science Society of America Proceedings 39, 377–379. L’Annunziata, M.F. (1984) Agricultural biochemistry: reaction mechanisms and pathways in biosynthesis. In: L’Annunziata, M.F. and Legg, J.O. (eds) Isotopes and Radiation in Agricultural Sciences, Vol. 2: Animals, Plants, Food and the Environment. Academic Press, London, pp. 105–182. L’Annunziata, M.F. (2003) Flow scintillation analysis. In: L’Annunziata, M.F. (ed.) Handbook of Radioactivity Analysis, 2nd edn. Elsevier, Amsterdam, The Netherlands, pp. 989–1062. L’Annunziata, M.F. and Fuller, W.H. (1971a) Soil and plant relationships of inositol phosphate stereoisomers; the identification of D-chiro- and muco-inositol phosphates in a desert soil and plant system. Soil Science Society of America Proceedings 35, 587–595. L’Annunziata, M.F. and Fuller, W.H. (1971b) Nuclear magnetic resonance spectra of acetate derivatives of soil and plant inositol phosphates. Soil Science Society of America Proceedings 35, 655–658. L’Annunziata, M.F. and Fuller, W.H. (1976) Evaluation of the mass spectral analysis of soil inositol, inositol phosphates, and related compounds. Soil Science Society of America Journal 40, 672–678. L’Annunziata, M.F. and Gonzalez, J. (1977) Soil metabolic transformations of carbon-14-myo-inositol, carbon-14phytic acid and carbon-14-iron(III) phytate. In: Soil Organic Matter Studies, Vol. 1. International Atomic Energy Agency, Publication No. IAEA-SM-211/66, Vienna, Austria, pp. 239–253. L’Annunziata, M.F., Fuller, W.H. and Brantley, D.S. (1972) D-chiro-Inositol phosphate in a forest soil. Soil Science Society of America Proceedings 36, 183–184. L’Annunziata, M.F., Gonzalez, J. and Olivares, L.A. (1977) Microbial epimerization of myo-inositol to chiro-inositol in soil. Soil Science Society of America Journal 41, 733–736. Lambert, J.B., Shurvell, H.F., Lightner, D.A. and Cooks, R.G. (1998) Organic Structural Spectroscopy. Prentice-Hall, Upper Saddle River, New Jersey. Nakajima, M., Tomida, I., Kurihara, N. and Takei, S. (1959) Zur Chemie des Benzolglykols. V. Eine neue Synthesis der Inositole. Chemische Berichte 92, 173–178. Peterbauer, T. and Richter, A. (1998) Galactosylinositol and stachyose synthesis in seeds of Adzuki bean. Plant Physiology 117, 165–172. Posternak, T. (1962) Scyllo-Inosose (myo-inosose-2): Bacterial oxidation of myo-inositol. In: Whistler, R.L., Wolfram, M.L., Bemiller, J.N. and Shafizadeh, F. (eds) Methods in Carbohydrate Chemistry. Academic Press, New York, p. 294.

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Reed, R.I., Reid, W.K. and Wilson, J.M. (1962) The mass spectra of some flavones and carbohydrates. In: Advances in Mass Spectrometry, Vol. 2. Pergamon Press, New York, pp. 416–427. Richter, A. Thonke, B. and Popp, M. (1990) D-1-O-methyl-muco-Inositol in Vicum album and members of the Rhizophoraceae. Phytochemistry 29, 1785–1786. Scholda, R., Billek, G. and Hoffmann-Ostenhof, O. (1964) Biosynthesis of cyclitols. VIII. Mechanism of the conversion of meso-inositol to D-pinitol and D-inositol in Trifolium incarnatum. Monatshefte für Chemie 95, 1311–1317. Smith, R.M., Chienthavorn, O., Wilson, I.D., Wright, B. and Taylor, S.D. (1999) Superheated heavy water as the effluent for HPLC–NMR and HPLC–NMR–MS of model drugs. Analytical Chemistry 71, 4493–4497. Sweeney, D.J., Lynch, G., Bidgood, A.M., Lew, W., Wang, K.-Y. and Cundy, K.C. (2000) Metabolism of the influenza neuraminidase inhibitor prodrug oseltamivir in the rat. Drug Metabolism and Disposition 28, 737–741. Turner, B.L., Papházy, M.J., Haygarth, P.M. and McKelvie, I.D. (2002) Inositol phosphates in the environment. Philosophical Transactions of the Royal Society, London, Series B 357, 449–469. Wanek, W. and Richter, A. (1995) Purification and characterization of myo-inositol 6-O-methyltransferase from Vigna umbellate Ohwi et Ohashi. Planta 197, 427–434.

5

Isolation and Assessment of Microorganisms That Utilize Phytate Jane E. Hill1 and Alan E. Richardson2 1

Environmental Engineering Program, Yale University, 9 Hillhouse Avenue, PO Box 8286, New Haven, CT 06520, USA; 2CSIRO, Plant Industry, PO Box 1600, Canberra, ACT 2601, Australia

The first phytase was discovered in the earlier part of the last century (Suzuki and Takaishi, 1907; Dox and Golden, 1911), but it was not until the efforts of Cosgrove and others in the 1960s (Cosgrove, 1980) that interest in identifying, characterizing and commercializing phytate-degrading organisms and their enzymes intensified. Indeed, it is becoming increasingly apparent that a range of microorganisms possess the ability to utilize phytate in their environment (Shieh and Ware, 1968; Irving and Cosgrove, 1971; Power and Jagannathan, 1982; Richardson and Hadobas, 1997; Yanke et al., 1998). These microorganisms occur in a variety of diverse habitats and do not employ an identical method to degrade phytate. Phytases, defined as enzymes that initiate the cleavage of myo-inositol hexakisphosphate, are now classified into four groups based on common biochemical and catalytic mechanisms. These are: (i) histidine acid phosphatases, which are widespread in fungi (Ullah et al., 1991; Van Etten et al., 1991); (ii) cysteine phosphatases, which have been identified in rumen bacteria (Yanke et al., 1999; Chu et al., 2004); (iii) β-propeller phytases, which occur mainly in the bacilli bacteria (Kim et al., 1998b; Ha et al., 1999; Shin et al., 2001; Oh et al., 2004); and (iv) purple acid phosphatases, which occur in a variety of organisms. The phytases are described in detail elsewhere in this volume (see Mullaney and Ullah, Chapter 7). The existence of phytate-degrading microorganisms challenges the conventional perception

that inositol phosphates are recalcitrant in the environment and of limited biological availability. This has arisen from the strong stabilization of inositol phosphates following sorption to clays or precipitation with metals (see Celi and Barberis, Chapter 13, this volume), their abundance in soils (see Turner, Chapter 12, this volume) and their limited availability to plants (see Richardson et al., Chapter 15, this volume). Our understanding of phytate-degrading microorganisms has been limited by a lack of ecological information and the difficulty in isolating and assessing the causative microorganisms in vivo. This chapter describes procedures for screening phytate-degrading microorganisms and characterizing their phytase activity, and provides recommendations for future studies. The major groups of microorganisms that have been isolated and characterized are summarized and current ecological information for these organisms is discussed.

Assessing Microorganisms for Phytate-degrading Activity Microorganisms able to degrade phytate have been isolated from a range of terrestrial and aquatic environments and their optimal cultivation conditions are often genera-specific. They include yeasts, filamentous fungi and Grampositive as well as Gram-negative bacteria. Three

©CAB International 2007. Inositol Phosphates: Linking Agriculture and the Environment (eds B.L. Turner, A.E. Richardson and E.J. Mullaney)

61

J.E. Hill and A.E. Richardson

microorganisms, in particular Aspergillus niger, Escherichia coli and Peniophora lycii, have been studied in detail and are produced commercially as an additive for animal feeds (Simon and Igbasan, 2002). However, the inherent variety in phytatedegrading microorganisms and the phytases that they produce suggests that other types of organisms and phytases await characterization. The diversity of phytate-degrading microorganisms is immense and requires a more detailed categorization than that used for the enzymes themselves. Detailed information on phytatedegrading microorganisms and the ecosystems in which they are found is fundamental to improving our understanding of inositol phosphates in the environment.

Screening for phytate utilization The most common procedure for screening microorganisms for potential to degrade phytate involves growth on solid laboratory media containing sodium- or calcium-phytate (Shieh and Ware, 1968; Cosgrove et al., 1970; Richardson and Hadobas, 1997). Calcium-phytate forms a precipitate at the pH range used in most media (pH 5–8) and can indicate the presence of phytate-degrading organisms in two ways. First, a microorganism growing on phytate as the sole source of phosphorus should only be able to do so (a)

(b)

pH 5

8.0

8

6.0

6

4.0

4

2.0

2

0

0 12

24

36

80

)

10

60

(

pH 7

10.0

40 20

Phosphate released (% total) ( )

Calcium-phytate Sodium-phytate content (mg/ml)

Sodium-phytate

due to its ability to release an extracellular phytase or by uptake through the outer membrane and its subsequent dephosphorylation within the periplasm (see Greiner, Chapter 6, this volume). Second, a clear zone around colonies that grow on calcium-phytate media indicates capacity to solubilize phytate presumably through the production of acids or chelates (Richardson and Hadobas, 1997; Mukesh et al., 2004; Fig. 5.1). Appropriate controls must be included to show that isolates do not grow when presented with a medium with all the ingredients except phytate. False positives may result from trace amounts of non-phytate phosphorus being present in the inocula or medium ingredients. A diagnostically false zone of clearing around a colony may be caused by the production of acid alone. Not all organisms that grow on solid media will grow easily in liquid culture (Richardson and Hadobas, 1997), so isolates can also be assessed for phytate utilization using liquid media. Growth of an organism in liquid culture containing phytate as the sole source of phosphorus should, with correct controls, provide a better indication of the ability of an organism to degrade phytate. Care must be taken to ascertain that the reason for growth is the degradation of phytate. Adequate controls showing a lack of growth as well as the removal of trace amounts of inorganic phosphate from the inocula are essential. Even with these precautions, growth may still occur (e.g. from phosphorus derived from

Bacterial growth (log10 cells/ml) ( )

62

0

48

Time (h)

Fig. 5.1. Screening for phytate-degrading microorganisms. (a) Growth of Pseudomonas sp. strain CCAR59 on agar containing sodium-phytate and calcium-phytate as the sole source of carbon and phosphorus at pH 7 and 5. (b) Degradation of sodium-phytate and release of phosphate to the culture supernatant during growth (pH 5.5) of strain CCAR59 in liquid culture. (From Richardson and Hadobas, 1997.)

Microorganisms That Utilize Phytate

lysis of the inoculum cells). Such procedures, therefore, only indicate the potential to degrade phytate and should not be used in isolation. Both solid and liquid screening procedures do not confirm phytase activity, and characterization of phytase activity and/or enzyme purification is necessary, as outlined below. It should also be considered whether microorganisms are using phytate as a source of carbon, phosphorus or both. While some cultured organisms can survive on phytate as a sole source of carbon and phosphorus (e.g. Pseudomonas spp. and Klebsiella aerogenes; Irving and Cosgrove, 1971; Tambe et al., 1994), other organisms may not (Richardson and Hadobas, 1997). A further important point is the nature of the carbon source used. Phosphorus bioavailability is often dependent on the ability of an organism to solubilize the source of phosphorus. Substrates such as citrate, which is a chelating agent, may make phytate more bioavailable by influencing substrate solubility (Hernandez et al., 2003; Munoz and Valiente, 2005).

Case study of the isolation of phytatedegrading Pseudomonas spp. from soil Two strategies to isolate phytate-degrading microorganisms were used by Richardson and Hadobas (1997). In one approach, agar medium containing sodium-phytate was used to identify microorganisms with the potential to degrade phytate. This indicated that 63% of isolates were able to grow on the defined medium containing phytate as the sole source of phosphorus and carbon, relative to tryptic-soy agar (a general growth medium for heterotrophic bacteria). When a 100member subset of these microorganisms was grown in liquid culture of the same composition, none were able to grow. However, 39% of the isolates were able to grow when citrate was added to the medium, although none were able to liberate phosphate from phytate as determined by the presence of molybdate-reactive phosphate in the culture supernatant. The second approach used enrichment culture and was more successful in identifying isolates that could degrade phytate. Liquid cultures contained phytate as the sole source of phosphorus either with or without the addition of citrate as a carbon source (Richardson and Hadobas,

63

1997). After purification of broth isolates by agar medium of the same composition, 44% of isolates were able to grow when reinoculated into the original broth medium. Of these, 4% liberated phosphate in the culture supernatant, from which six genetically unique bacterial strains were isolated. These isolates showed greatest similarity to Pseudomonas spp., which are a ubiquitous group of microorganisms found in soil. Growth of one of these isolates (strain CCAR59) on sodiumphytate and calcium-phytate media as the sole source of carbon and phosphorus is shown in Fig. 5.1, with solubilization of calcium-phytate being evident at pH 5. This example highlights the importance of using a combination of growth conditions and assay procedures to verify that microorganisms are able to utilize phytate.

Characterization of phytase activity Following the isolation of microorganisms that have the ability to degrade phytate in laboratory media it is necessary to confirm phytase activity. Ideally this is achieved through purification of the enzyme and characterization of its kinetics, although this is often a challenging and timeconsuming task. First, organisms must be grown in culture to generate a sufficient amount of enzyme to enable purification. Then, a series of purification steps, such as ammonium sulphate fractionation and chromatographic separation, are used to purify the enzyme. Molecular weight, isoelectric point, specific activity, thermal stability and pH optima can then be evaluated (e.g. Greiner et al., 1993; Shimizu, 1993; Greiner et al., 1997; Choi et al., 2001). Colorimetric assays, which measure free phosphate in solution, are useful for assessing phytase activity. Typically, microbial extracts or extracellular components are incubated in the presence of sodium-phytate under defined conditions (i.e. temperature, pH, etc.) and released phosphate is determined at intervals. This approach is primarily used for initial assessment of phytase activity from different microorganisms, subsequent tracking of enzymes during purification procedures, determination of kinetics and establishment of an unequivocal link between biological synthesis of the enzyme and the utilization of phytate.

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J.E. Hill and A.E. Richardson

Antibody-based technologies have also been used to track purification as well as to localize phytase in microorganisms and tissues (Ullah and Gibson, 1987; Golovan et al., 2001). Immunoassays use antibodies to recognize highly specific motifs (generally protein domains) that are generated from the immune response of animals. Mono- and polyclonal antibodies have been raised for phytases from A. niger (Ullah and Gibson, 1987; Hostetler et al., 2005) and E. coli (Golovan et al., 2001).

Case study of phytase purification and characterization from Klebsiella terrigena Klebsiella terrigena is a soil bacterium that can utilize phytate as the sole source of phosphorus. Phytase production is greatest when the culture reaches the stationary phase and the activity of the enzyme is stimulated by the presence of phytate (see Greiner, Chapter 6, this volume). Greiner et al. (1997) purified the enzyme, characterized its substrate specificity and kinetics, and determined the products of enzymatic hydrolysis using high-performance liquid chromatography (HPLC). Protein purification to this extent (i.e. as outlined in Table 5.1) is necessary to characterize the enzyme. Preparation purity is ascertained using sodium dodecyl sulphfate–polyacrylamide gel electrophoresis (SDS– PAGE), which can also be used to indicate protein

molecular mass and the extent of glycosylation. Once purified, phytases from different sources can be compared (Table 5.2).

Assessment of phytase activity by analytical methods Major techniques that have been applied for assessing the catalytic function of phytases from different microorganisms include ion chromatography, HPLC and 31 P nuclear magnetic resonance (NMR) spectroscopy. These techniques are particularly useful for investigating the presence of lower-order inositol phosphates and inorganic phosphates to establish pathways of degradation (Freund et al., 1992; Skoglund et al., 1997; Cade-Menun, 2005). For example, Greiner et al. (1997) used ion pair chromatography to conduct a time course study of the degradation of sodium-phytate by K. terrigena phytase (Fig. 5.2). The results showed the accumulation of an inositol trisphosphate end product, although the presence of positional isomers could not be determined. The dephosphorylation pathway for several Bacillus spp. that degrade phytate has similarly been determined (Greiner et al., 2002). In this case, the genes responsible for phytate degradation from Bacillus subtilis 168, B. amyloliquefaciens ATCC 15841 and B. amyloliquefaciens 45 were

Table 5.1. Purification of phytase from Klebsiella terrigena showing higher specific activity with increasing purity. (From Greiner et al., 1997.) Purification step Crude extract (NH4)2SO4 precipitation CM-Sepharosed DEAE-Sepharosee Mono Sf Sephacryl S-200g

Total activitya (U)

Specific activityb (U/mg)

Recoveryc (%)

268.0 255.0 178.0 112.0 90.0 76.0

0.5 1.5 55.6 80.0 102.3 205.0

– 95 66 42 34 28

Total activity (U) is determined where 1 unit liberates 1 µmol phosphate per minute. Specific activity is the total activity divided by total protein in the sample. c Recovery is the total activity in the step divided by the total activity of the crude extract expressed as a percentage. d CM-Sepharose is the cationic exchange resin carboxymethyl-Sepharose. e DEAE-Sepharose is the anionic exchange resin diethylaminoethyl-Sepharose. f Mono S is a cation exchange resin. g Sephacryl S-200 is a hydrophilic resin. a b

Microorganisms That Utilize Phytate

Table 5.2. Kinetics constants (kcat) for degradation of myo-inositol hexakisphosphate and its lowerorder esters by phytase from Bacillus amyloliquefaciens (strain ATCC 15841). (From Greiner et al., 2002.) Substrate

kcat (/s)

myo-Inositol hexakisphosphate D-myo-Inositol 1,2,4,5,6pentakisphosphate D-myo-Inositol 1,2,3,5,6pentakisphosphate D-myo-Inositol 1,2,3,4,5pentakisphosphate D-myo-Inositol 1,2,5,6tetrakisphosphate D-myo-Inositol 1,2,6-trisphosphate myo-Inositol 2-monophosphate

18.5ca 16.0a 7.3b 7.5b 10.1d 1.9e 0.12g

a

Means accompanied by a different letter are significantly different (P < 0.05).

Inositol phosphates (µg)

cloned and expressed in a common strain of bacilli (B. subtilis MU331), and were compared to phytases from E. coli, A. niger and rye (Secale cereale L.). Degradation products were determined by HPLC and demonstrated that Bacillus spp. had two independent pathways for degradation of phytate. This initially led to an accumulation of either D/L-myo-inositol 2,4,5-trisphosphate or myo-inositol 2,4,6-trisphosphate and subsequently to the generation of myo-inositol 2-monophosphate (for further details see Greiner, Chapter 6, this volume).

40

IP6

30 20

IP4

IP5

10

IP3

65

Sources of Phytase and Expression in Microorganisms Phytate-degrading microorganisms Phytate-degrading microorganisms have been isolated from a wide range of environments, including marine and fresh water ecosystems, soils, sediments and the gastrointestinal tract of animals (Table 5.3). Although no systematic study has been conducted across different environments, it is evident that the phytase phenotype is manifest in a variety of habitats. This includes a variety of soils (Richardson and Hadobas, 1997), organicrich and organic-poor locations (Choi et al., 2001; Kim et al., 2003) and anaerobic environments (Yanke et al., 1999). The anaerobic environment is of particular interest, because inositol phosphates appear to be degraded rapidly under anaerobic conditions (Suzumura and Kamatani, 1995a) and are absent from submerged wetland soils where anaerobicity is common (Turner and Newman, 2005). Anaerobic environments may therefore be important in the inositol phosphate cycle and an important source of novel phytate-degrading microorganisms. The mixture of intracellular and extracellular enzymes (Table 5.3) is also significant and suggests that microorganisms use different mechanisms to hydrolyse inositol phosphates, either in the external environment or within the periplasm. Extracellular enzymes are generally more tolerant to pH and temperature fluctuation and show greater resistance to proteolytic degradation (see George et al., Chapter 14, this volume). These traits will not only affect the persistence and effectiveness of phytase from different microorganisms in different environments, but are also attractive features for the commercial development of phytases.

0 0

120

240

360

480

Time (min)

Fig. 5.2. Time course for the degradation of sodium-phytate by Klebsiella terrigena phytase as determined by ion pair chromatography. (From Greiner et al., 1997 with permission from Elsevier.) myo-Inositol hexakisphosphate (IP6) is hydrolysed to a myo-inositol trisphosphate (IP3) end product. IP4, myo-inositol tetrakisphosphate; IP5, myo-inositol pentakisphosphate.

Production of microbial phytases Both native and recombinant phytase enzymes have been assessed in an effort to optimize production. Filamentous fungi in particular have been utilized extensively for their high production efficacy. However, E. coli and yeast (Saccharomyces cerevisiae) (Phillippy and Mullaney,

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J.E. Hill and A.E. Richardson

Table 5.3. Source of phytate-degrading microorganisms, their environment of isolation, oxygen requirement and location of enzyme expression. Oxygen requirementa

Enzyme locationb

Reference

A A A A

EX EX IN EX

Choi et al. (2001) Kim et al. (1998a) Kim et al. (2003) Yoon et al. (1996)

A

IN

Italian sourdough

A

IN

Cellulosic waste

A

EX

Jareonkitmongkol et al. (1997) De Angelis et al. (2003) Mitchell et al. (1997)

Soil Cattle farm soil

A A

EX IN

Tseng et al. (2000) Cho et al. (2003)

Soil

A

IN

Laboratory stock (fresh water organism) Dried flower buds

A

IN

Richardson and Hadobas (1997) Freund et al. (1992)

A

IN

Tempehc

A

IN/EX

Rumen fluid

AN

EX

Microorganism

Environment

Bacillus sp. KHU-10 Bacillus sp. DS11 Citrobacter braakii YH-15 Enterobacter sp. 4

Boiled rice Cattle shed floor Sea water Soil of leguminous plants Soil

Klebsiella oxytoca MO-3 Lactobacillus sanfranciscensis CB1 Myceliophthora thermophila (48102) Penicillium simplicissimum Pseudomonas syringae MOK1 Pseudomonas spp. Paramecium tetraurelia 51s Pichia anomala

Rhizopus oligosporus CT11K2 Selenomona ruminatium

Vohra and Satyanarayana (2002) Sutardi and Buckle (1988) Yanke et al. (1999)

a

A/AN refers to aerobic or anaerobic production, respectively. Expression is either intracellular (IN) or extracellular (EX). c A fermented food typically made from soybean that is popular in South-east Asia. b

1997; Han et al., 1999; Kerovuo et al., 2000; Sajidan et al., 2004) and plant systems have also been employed (Pen et al., 1993; Li et al., 1997; Kusnaki et al., 1998; Ponstein et al., 2002). There is still much to be understood about the basic stimuli and repression systems that are associated with the expression of phytases in native systems. However, phosphorus limitation, in the form of either a low intracellular or ambient phosphorus concentration, is a key stimulus for phytase production in many microorganisms. This implies that phosphorus limitation is paramount to the expression of genes for phytase synthesis in microorganisms. However, there are notable exceptions to this: e.g. the rumen bacteria reported by Yanke et al. (1998), which live in an environment rich in phosphorus.

Optimization of cultures for expression of phytase Growth conditions significantly affect the expression and production of phytase, but no generic methodology exists to assess this and few trends are apparent. E. coli has two periplasmic phosphatases that can accept phytate as a substrate. The first, agp, has a broad substrate range, is constitutively expressed and may be used to scavenge glucose from glucose 1-phosphate (Wanner, 1990). The second, appA, has a narrow substrate range and its expression (and production) is stimulated by entry into the stationary phase of growth, a low ambient phosphate concentration and anaerobic conditions (Touati et al., 1987; Greiner et al., 1993; see Greiner, Chapter 6, this volume).

Microorganisms That Utilize Phytate

Induction of the PHO regulon is not implicated in the synthesis of the enzyme (Touati et al., 1987) nor is it induced by phytate (Greiner et al., 1993). Under aerobic growth conditions, growth restriction by low inputs of carbon, nitrogen and sulphur had no influence on the production of either of the E. coli enzymes (Touati et al., 1987). Bacillus spp. (strain KHU-10) produce a high level of extracellular phytase in the stationary phase when grown in complex media containing maltose, peptone and beef extract (Choi et al., 1999). Stimulation of phytase production in B. subtilis has also been observed by the addition of phytate to the cultivation medium (Kim et al., 1999b). In more simple but defined media, Bacillus phytases require calcium ions for effective phytase activity. However, this requirement appears to influence only the stability of the enzyme rather than regulate its production (Shimizu, 1992; Choi et al., 1999). In more complex media (e.g. wheat bran medium), factors that contribute to high phytase production are not well characterized, but their impact on production and activity of phytase is significant (Shimizu, 1992; Choi et al., 1999). The low solubility of phytate in wheat bran is thought to result in a controlled release of substrate, which directly regulates phosphate in the medium and, therefore, enzyme production. However, other factors in the medium may also be important. Production of phytase in fungi is typically enhanced by low phosphate in the medium and is repressed at higher concentrations (Shieh and Ware, 1968; Han and Gallagher, 1987; Vats and Banerjee, 2002; Kim et al., 2003). Mutagenesis of cultures (e.g. through irradiation) results in increased phytase production: e.g. in A. niger NRRL3135 (Chelius and Wodzinski, 1994). However, a decrease in enzyme production occurs when the medium contains a high concentration of glucose and/or is poorly aerated. Saccharomyces cerevisiae produces three extracellular phosphatases (PHO5, PHO10 and PHO11) that can hydrolyse phytate as well as a range of other phosphate monoesters (Nakamura et al., 2000; Andlid et al., 2004). These genes are associated with the PHO regulon and respond to low levels of extracellular phosphate (Lemire et al., 1985) in a complex regulatory framework (Ogawa et al., 1993; Kaffman et al., 1994; see Greiner, Chapter 6, this volume).

67

Production of phytases on a large scale has also been undertaken using submerged and solidstate fermentation technologies. As with lab-scale cultures, each strain has optimal conditions, including the rate of aeration, temperature, inoculum conditions and media components. These have been best studied for A. niger NRRL3135 (Howson and Davis, 1983; Han and Gallagher, 1987; Ullah and Gibson, 1987; Ebune et al., 1995; Krishna and Nokes, 2001). Submerged fermentation studies for E. coli and Bacillus sp. DS11 show that phytase production was essentially the same as for smaller-scale culture studies (Kim et al., 1998a; Kleist et al., 2003). Large-scale production is an essential subsequent step for the commercialization of enzymes. Generally this is best achieved when phytaseencoding genes are cloned and expressed in recombinant microorganisms using expression vectors, where protein production can more easily be optimized and yields are often substantially higher (Quax, 1997; Han et al., 1999; Kim et al., 1999a; Rodriguez et al., 2000a; Xiong et al., 2004; Table 5.4). In contrast, optimization of culture conditions for the expression of ‘native’ enzymes is more difficult. Observations based on composition of the media are often inconclusive, due in large part to the complexity of the regulation of phytase gene expression and conditions for enzyme activation in diverse microorganisms. These difficulties notwithstanding, culture optimization can provide valuable insight into the regulation of the enzyme in vivo.

Properties of phytases from microorganisms Catalytic and physicochemical properties of phytases from some representative microorganisms are shown in Table 5.5. Phytase from Citrobacter braakii has the highest reported specific activity (Kim et al., 2003). The E. coli appA phytase with an optimal pH 2.5 has the highest reported kcat (Dassa et al., 1982; Greiner et al., 1993) and has been subjected to a number of mutagenic modifications (Golovan et al., 2000; Rodriguez et al., 2000b). Similarly, phytase (phyA) from A. niger NRRL 3135 (formerly A. ficuum) has been extensively studied in terms of catalytic and physical

68

J.E. Hill and A.E. Richardson

Table 5.4. Host expression systems and cultural conditions for expression of phytases from diverse microorganisms. Microorganism and source of phytase

Expression host

Aspergillus niger

E. coli

5.1

43–50

Saccharomyces cerevisiae Pichia pastoris A. niger (native)

2.0–2.5, 5.0–5.5

55–60

2.5, 5.5 2.2, 5.0–5.5

60 55–58

E. coli P. pastoris

4.5 3.5

55 60

Pseudomonas putida E. coli (native) B. subtilis B. subtilis 168 (native) B. subtilis B. licheniformis (native) E. coli Bacillus sp. DS11 (native)

4.0

55

4.5 7.0 6.0–7.5

55–60 55 55–60

7.0 6.0–7.5

65 55–60

– 7.0

70 70

Escherichia coli

Bacillus subtilis 168

B. licheniformis

Bacillus sp. DS11

pH optima

properties and mutational analysis (e.g. Wyss et al., 1999a,b; Lehmann et al., 2002; Tomschy et al., 2002). A. niger, with two pH optima (2.2 and 5.0–5.5; Ullah and Gibson, 1987), is a particularly useful enzyme due to its ability to function effectively under gastrointestinal conditions relevant for swine and poultry. The cloning and expression of this phytase (Mullaney et al., 1991; van Hartingsveldt et al., 1993) has led to its commercial development as a feed additive (e.g. Natophos®). Likewise the phytases from P. lycii (Lassen et al., 2001) and E. coli (Fairley, 1998) have been developed commercially. From a commercial perspective, phytases destined for addition to the diet of animals need to meet a number of specific requirements. Animal feeds are often produced through a pelleting process at temperatures between 65°C and 80°C. Thermostability of the phytase is therefore desirable. Phytases are also required to function within the gastrointestinal tract of various animals that have unique body temperature and digestive pH (Riley and Austic, 1984; Radcliffe et al., 1998;

Temperature optima (ºC)

Reference Phillippy and Mullaney (1997) Han et al. (1999) Han and Lei (1999) Ullah and Gibson (1987), Wyss et al. (1999a,b) Golovan et al. (2000) Rodriguez et al. (2000a) Dharmsthiti et al. (2005) Greiner et al. (1993) Tye et al. (2002) Shimizu (1992), Kerovuo et al. (1998) Tye et al. (2002) Shimizu (1992), Kerovuo et al. (1998) Kim et al. (1998b) Kim et al. (1998a)

Ramseyer et al., 1999). For example, the adult body temperature for swine is around 39°C, whereas it is between 5ºC and 18°C in fish. Agedependent variations must also be considered and resistance of phytases to proteolytic cleavage is important (Wyss et al., 1999b). Modification of phytases to meet such requirements has been studied extensively (e.g. Wyss et al., 1999a,b; Lehmann et al., 2002; Tomschy et al., 2002).

The Ecology of Phytate-degrading Microorganisms Inositol phosphates exist in a wide range of environments including arable, forest and grassland soils, as well as fresh water and marine sediments (Turner et al., 2002; see Turner, Chapter 12, and McKelvie, Chapter 16, this volume). Given that productivity in many of these ecosystems can be limited by the availability of phosphorus, why do inositol phosphates persist when microorganisms are present that can potentially utilize them?

Table 5.5. Catalytic and physicochemical properties of phytases from diverse microorganisms (values denoted by a dash are not available).

Microorganism

pI 4.5–5.2

pH optima

5.0

5.0–5.5

70

Bacillus licheniformis B. subtilis

5.0 6.3–6.5

4.5–6.0 6.0–7.5

Bacillus sp. DS11 Citrobacter braakii Escherichia coli Lactobacillus sanfranciscensis Klebsiella terrigena Penicillium simplicissimum Peniophora lycii Pseudomonas syringae MOK1 Saccharomyces cerevisiae Schwanniomyces castellii

5.3 – 6.0–7.4 5.0

5.8 3.6 –

Specific activitya (U/mg)

55–58

k m (µmol)

Substrate range

Reference

50–103

10–40

Narrow

142–196

11–23

Narrow

55–60 55–60

– 9–15

– 50–500

Narrow Narrow

7.0 4.0 4.5 4.0

70 50 55–60 45

– 3457 811–1800 6–73

550 460 130–630 –

Narrow Narrow Narrow Middle

Ullah and Gibson (1987), Wyss et al. (1999a) Yamada et al. (1968), Wyss et al. (1999a,b) Tye et al. (2002) Shimizu (1992), Kerovuo et al. (1998) Kim et al. (1998a) Kim et al. (2003) Greiner et al. (1993) De Angelis et al. (2003)

5.0–6.0 4.0 4.0–4.5 5.5

55 55 45 40

205 3 – –

300 – – 380

Middle Broad Narrow Narrow

Greiner et al. (1997) Tseng et al. (2000) Lassen et al. (2001) Cho et al. (2005)

2.0–2.5, 5.0–5.5 4.4

55–60 77

– 418

– 38

Broad Broad

Han et al. (1999) Segueilha et al. (1992)

Microorganisms That Utilize Phytate

Aspergillus niger NRRL3135 A. terreus

– –

2.5, 5.0–5.5

Temperature optima (ºC)

a

Determined at 37ºC, with the exception of Schwanniomyces castellii, which was determined at 70ºC.

69

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J.E. Hill and A.E. Richardson

Perhaps the most important factor that regulates the availability of inositol phosphates to microorganisms is the formation of strong and stable complexes between inositol phosphates and metals such as iron, aluminium, copper, calcium and magnesium (Vohra et al., 1965; see Celi and Barberis, Chapter 13, this volume). Persistence of inositol phosphate in soils therefore may be linked to the ability of microorganisms to solubilize and access these forms of phosphorus. Alternatively, many soils do not represent phosphorus-limited environments for microorganisms (Jakobsen et al., 2005), reducing the need for microorganisms to utilize inositol phosphates. In this scenario, inositol phosphate concentrations would be linked to the degree of phosphorus limitation, and there is tentative evidence for this in some soils (see Turner, Chapter 12, this volume). To date much of our understanding of phytase from microorganisms has been driven by the potential for commercial phytase production, rather than to obtain an ecological understanding. Clearly, there is a need to better understand the ecological importance of microorganisms that have the potential to degrade phytate in different ecosystems.

Function of phytases in different ecosystems Plate and liquid culture screening assays are a simple and effective means to obtain general information on potential phytate-degrading activity of microorganisms in environmental samples. However, as described above, such assessment may result in erroneous estimates of the number of organisms with phytate-degrading capability (Richardson and Hadobas, 1997; Fig. 5.1). For example, Greaves and Webley (1965) estimated soil and rhizosphere populations of phytatedegrading bacteria to constitute between 30% and 48% of the total population using plate-based screening methods. While this may be an overestimate, Unno et al. (2005) recently isolated and characterized a large number of microorganisms, predominantly Burkholderia spp., from the rhizosheath and rhizoplane of lupin (Lupinus albus L.) plants. Interestingly, many of these isolates were also able to promote plant growth, suggesting the importance of phytate hydrolysis in close proximity to plant roots (see Richardson et al., Chapter

15, this volume). Although such screening procedures do not unequivocally demonstrate the function of phytate-degrading microorganisms in soil, they do indicate a potential role. Further work is needed to establish the function of phytatedegrading microorganisms in such environments. An important advance in this area is the development of a substrate analogue for phytate which would offer the possibility of rapid screening procedures (Berry and Berry, 2005). An interesting ecosystem that needs to be more thoroughly investigated is the gastrointestinal tract of animals. Yanke et al. (1998, 1999) reported the isolation of anaerobic microorganisms (e.g. Selenomona ruminatium) from the rumen fluid of cows and suggested their role in phytate utilization. There is evidence that metal complexation in cattle fed a grain-based diet can result in manures that contain appreciable concentrations of inositol phosphates (see Dao, Chapter 11, this volume). However, it is generally considered that cattle do not excrete significant amounts of inositol phosphates (see Leytem and Maguire, Chapter 10, this volume). Possible explanations are that ruminants contain a pH-neutral foregut that is effective for phytate degradation, or that hydrolysis of inositol phosphates occurs under anaerobic–fermentative conditions. Plant-derived phytases in feed may also be active in the rumen (see Lei and Porres, Chapter 9, this volume). Monogastric animals, on the other hand, have fermentative digestion after an acidic stomach, which may either be detrimental to incoming phytate-degrading bacteria or denature phytases. Thus, poultry manures tend to contain large amounts of phytate (e.g. McGrath et al., 2005). In spite of this, there is some evidence indicating that bacteria in the hindgut of monogastric animals might also hydrolyse phytate, resulting in a decrease in the excreta of swine and poultry (Leytem et al., 2004). Further work to understand the degradation of phytate in the gut of both monogastric animals and ruminants is required.

Ecological understanding of phytatedegrading microorganisms In understanding the potential for degradation of inositol phosphates in terrestrial and aquatic systems, knowledge of the geochemistry of the environment is required. Microorganisms live in niche

Microorganisms That Utilize Phytate

environments, so any gross parameters measured should be considered in this context. Interaction between microorganisms and inositol phosphates requires that the substrate be available to the microorganism, although this can be influenced by a range of physiological and abiotic processes. Factors affecting the solubility of inositol phosphates in soil include pH, clay content, and the presence and concentration of metal ions in soil solution (reviewed by Turner et al., 2002; see Turner, Chapter 12, and Celi and Barberis, Chapter 13, this volume). In particular, complexation of inositol phosphates with metals such as iron and aluminium can render the resulting insoluble phytates unavailable to microorganisms (e.g. Greenwood and Lewis, 1977). Microorganisms also require carbon and nitrogen and suitable physiological conditions such as pH and moisture, although phosphorus rarely limits microbial growth ( Jakobsen et al., 2005). An exception may be fresh water planktonic cells (Schindler, 1971; O’Sullivan, 1992; McComb and Davis, 1993), but in most soilbased environments the concentration of phosphate, or organic phosphates other than inositol phosphates, may provide sufficient phosphorus to support microbial growth. Molecular approaches to the ecology of phytate-degrading microorganisms Culture-independent approaches based on DNA technology indicate that generally <1% of microorganisms, particularly bacteria, can be cultured from environmental samples (Kaeberlein et al., 2002). Of those that have been cultured and characterized, few have been assessed for their ability to degrade phytate. It is therefore unclear whether the ability to utilize phytate is a common trait or restricted to select genera (e.g. as listed in Table 5.3). However, given the level of diversity present in phytatedegrading microorganisms so far isolated, and the vast number of microorganisms in environmental samples that have yet to be cultured, it seems likely that the number of microorganisms able to degrade phytate is large. Development of specific molecular tools to analyse environmental samples could be used to address this issue. Molecular community diversity analysis of ecosystems can be conducted using DNA amplification techniques based on the polymerase chain reaction (PCR). These are particularly useful when applied to the 16S ribosomal gene,

71

which is ubiquitous and has highly conserved sequence domains across the eubacterial and archaebacterial kingdoms. Techniques such as terminal-restriction fragment length polymorphism (T-RFLP) and denaturing gradient gel electrophoresis (DGGE) can be used to investigate the structure of microbial communities in environmental samples and identify ‘shifts’ in major groups of bacteria (Tajima et al., 2001; Possemiers et al., 2004; Van der Gucht et al., 2005). Such approaches could be applied to environment samples where concentrations of inositol phosphates are known to change (e.g. after the addition of manures to soil) to identify key groups of microorganisms that might be involved in their hydrolysis. A key challenge for future studies is to link changes in microbial community structure to specific functional groups of bacteria and/or specific biochemical processes within the environment. Molecular-based techniques can also be used to identify the presence (and ideally the function) of specific genes. To study gene shifts, sufficient DNA homology among target genes is required and at present this may be possible with the β-propeller phytases. This family of phytases has been shown (Cheng and Lim, 2006) to occur in a wide range of bacteria isolated from diverse environments (e.g. as identified using online data of the National Center for Biotechnology Information, available at www.ncbi.nlm.nih.gov). Preliminary work has shown that Shewanella oneidensis MR-1 phytase shares amino acid homology with a Shewanella organism in sea water samples from the Sargasso Sea (Venter et al., 2004). Based on this assessment and enzyme kinetics of the S. oneidensis phytase, it is possible that Shewanella plays a significant role in the degradation of phytate that has been observed in marine environments (Suzumura and Kamatani, 1995b; Cheng and Lim, 2006). An alternative approach is to use sets of β-propeller-specific nucleic acid primers to probe microbial communities and identify, and possibly isolate, microorganisms with homologous genes ( J. Hill, 2006, unpublished data).

Conclusions and Future Research It is becoming clear that microorganisms with the ability to utilize inositol phosphates are ubiquitous in both terrestrial and aquatic environments.

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J.E. Hill and A.E. Richardson

This challenges the conventional perception that inositol phosphates are recalcitrant and do not contribute greatly to the nutrition of organisms. The ecological implications of this abundance of phytate-degrading microorganisms remain largely unexplored, but are likely to be significant in several environments. For example, inositol phosphates can no longer be considered unimportant in terms of water quality (see McKelvie, Chapter 16, this volume). Assessment of phytate-degrading microorganisms requires screening (usually starting with solid or liquid enrichment), identification of the organism responsible, purification of the enzyme, and evaluation of growth parameters and enzyme properties. While current protocols are labourintensive, they provide important information concerning the potential of different groups of microorganisms to degrade phytate and, increasingly, of their ecological significance. However, much remains to be achieved. The use of higher throughput systems (e.g. robotic sampling and

culturing systems) should allow a wider diversity of environments to be investigated in detail. Irrespective of this, it is important that efforts to analyse microbial populations for phytate degradation also consider the environment in which the microorganisms operate. While the development of new screens that adopt techniques such as community and gene analysis will clearly assist in this process, these approaches must adopt meaningful ways to link the analysis of community structure with microbial function and ecosystem characteristics. Molecular approaches will need to incorporate analytical tools such as 31P NMR spectroscopy to determine the speciation and concentration of inositol phosphates. Uniting molecular and analytical tools is complex and will require collaborative expertise. However, until the development of more gene- and enzyme-specific approaches, these are important ways to gain an ecological understanding of phytate-degrading organisms and their role in the utilization of inositol phosphates in the environment.

References Andlid, T.A., Veide, J. and Sandberg, A.S. (2004) Metabolism of extracellular inositol hexaphosphate (phytate) by Saccharomyces cerevisiae. International Journal of Food Microbiology 97, 157–169. Berry, D.F. and Berry, D.A. (2005) Tethered phytic acid as a probe for measuring phytase activity. Bioorganic and Medicinal Chemistry Letters 15, 3157–3161. Cade-Menun, B. J. (2005) Characterizing phosphorus in environmental and agricultural samples by phosphorus-31 nuclear magnetic resonance spectroscopy. Talanta 66, 359–371. Chelius, M.K. and Wodzinski, R. J. (1994) Strain improvement of Aspergillus niger for phytase production. Applied Microbiology and Biotechnology 41, 79–83. Cheng, C. and Lim, B.L. (2006) Beta-propeller phytases in the aquatic environment. Archives of Microbiology 185, 1–13. Cho, J.S., Lee, C.W., Kang, S.H., Lee, J.C., Bok, J.D., Moon, Y.S., Lee, H.G., Kim, S. and Choi, Y. (2003) Purification and characterization of a phytase from Pseudomonas syringae MOK1. Current Microbiology 47, 290–294. Cho, J.S., Lee, C., Kang, S., Lee, J., Lee, H., Bok, J., Woo, J., Moon, Y. and Choi, Y. (2005) Molecular cloning of a phytase gene (phyM) from Pseudomonas syringae MOK1. Current Microbiology 51, 11–15. Choi, Y.M., Noh, D.O., Cho, S.H., Lee, H.K., Suh, H. J. and Chung, S.H. (1999) Isolation of a phytase-producing Bacillus sp, KHU-10 and its phytase production. Journal of Microbiology and Biotechnology 9, 223–226. Choi, Y.M., Suh, H. J. and Kim, J.M. (2001) Purification and properties of extracellular phytase from Bacillus sp KHU-10. Journal of Protein Chemistry 20, 287–292. Chu, H.M., Guo, R.T., Lin, T.W., Chou, C.C., Shr, H.L., Lai, H.L., Lai, H., Cheng, K., Selinger, B. and Wang, A. (2004) Structures of Selenomonas ruminantium phytase in complex with persulfated phytate: DSP phytase fold and mechanism for sequential substrate hydrolysis. Structure 12, 2015–2024. Cosgrove, D. J. (1980) Inositol Phosphates: Their Chemistry, Biochemistry and Physiology. Elsevier, Amsterdam, The Netherlands. Cosgrove, D. J., Irving, G.C. J. and Bromfield, S.M. (1970) Inositol phosphate phosphatases of microbiological origin: isolation of soil bacteria having inositol phosphate phosphatase activity. Australian Journal of Biological Sciences 23, 339–343.

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6

Phytate-degrading Enzymes: Regulation of Synthesis in Microorganisms and Plants Ralf Greiner

Federal Research Centre for Nutrition and Food, Centre for Molecular Biology, Haid-und-Neu-Strabe 9, D 76131 Karlsruhe, Germany

Phytate-degrading enzymes, also known as phytases, have a wide distribution in plants, microorganisms and some animal tissues (Konietzny and Greiner, 2002; see Hill and Richardson, Chapter 5, and Mullaney and Ullah, Chapter 7, this volume). They belong to a special class of phosphomonoesterases termed myo-inositol hexakisphosphate phosphohydolases, which are capable of initiating the stepwise release of phosphate residues from phytate (salts of myo-inositol hexakisphosphate), the major storage form of phosphate in plant seeds and pollen. The ability of such enzymes to hydrolyse phytate is usually known only from in vitro assays, and information on their in vivo function is rather limited. As enzymes are classified in general by their in vivo function, the term ‘phytate-degrading enzyme’ is preferred here to the term ‘phytase’, and is used throughout the chapter. The classification of enzymes as phytases based solely on in vitro assays becomes problematic when considering enzymes such as glucose-1-phosphatase in Escherichia coli and Enterobacter cloacae. This enzyme can hydrolyse phytate, albeit slowly, in vitro even though this is clearly not its in vivo function. Unless otherwise stated, designation of an enzyme as ‘phytatedegrading’ is based on in vitro assay using soluble sodium phytate.

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Phytate-degrading Enzymes and Their Classification Phosphomonoesterases are a diverse group of enzymes that encompass a range of sizes, structures and catalytic mechanisms. Based on the amino acid residue in the active site, phytatedegrading enzymes can be referred to as histidine acid phosphatases, β-propeller phosphatases, cysteine phosphatases and purple acid phosphatases (see Mullaney and Ullah, Chapter 7, this volume). Two classes of phytate-degrading enzymes are recognized by the International Union of Pure and Applied Chemistry and the International Union of Biochemistry (IUPAC–IUB): 3-phytase (EC 3.1.3.8), which initially removes phosphate from the D-3 position of the myo-inositol ring, and 6-phytase (EC 3.1.3.26), which preferentially initiates phytate dephosphorylation at the L-6 (D-4) position. However, phytate-degrading enzymes initiating phytate degradation at the D-5 and D-6 positions, respectively, have been found in nature (Barrientos et al., 1994; Greiner et al., 2000a). Phytate-degrading enzymes from microorganisms are considered to be 3-phytases, whereas 6-phytases are said to be characteristic of the seeds of higher plants. Most of the phytate-degrading enzymes studies so far with respect to their pathway of

©CAB International 2007. Inositol Phosphates: Linking Agriculture and the Environment (eds B.L. Turner, A.E. Richardson and E.J. Mullaney)

Synthesis in Microorganisms and Plants

phytate degradation fit into this consideration (Fig. 6.1). However, this is not a general rule, as exemplified by the indication of 3-phytase activity in lupine (Greiner et al., 2002) and soybean seeds (R. Greiner, 2000 unpublished data) and 6phytase activity in Paramecium (van der Kaay and van Haastert, 1995) as well as E. coli (Greiner et al., 2000a). However, it is worth mentioning that the 6-phytase activity of plant seeds initially hydrolyses the L-6 (D-4) phosphate residue from phytate, whereas phytate-degrading enzymes from Paramecium and E. coli initially remove the phosphate residue attached to the D-6 (L-4) position. Phytate-degrading enzymes are found in multiple forms, especially in plant seeds. These forms may even exhibit different stereospecificity of myo-inositol hexakisphosphate dephosphorylation, as reported recently for the phytate-degrading enzymes from lupines (Greiner et al., 2002). The phosphate residues of phytate are released by phytate-degrading enzymes at different rates and in different order. Independent of their bacterial, fungal or plant origin, the majority of the phytate-degrading enzymes exhibiting an optimum for phytate hydrolysis under acidic pH conditions release five of the six phosphate residues of phytate, and the final degradation product was identified as myo-inositol 2-phosphate (Konietzny and Greiner, 2002). Dephosphorylation of myoinositol 2-phosphate occurs only in the presence of high enzyme concentration during prolonged incubation. After removal of the first phosphate residue from phytate these histidine acid phytatedegrading enzymes continue dephosphorylation adjacent to a free hydroxyl group. The major phytate degradation pathways of 6-phytases of plant origin and 3-phytases differ only in the myoinositol pentakisphosphate intermediate generated (Fig. 6.1). The two exceptions reported so far are the 6-phytases from mung bean (Maiti et al., 1974) and wheat F2 phytase (Lim and Tate, 1973). The microbial 6-phytases, however, generate a completely different set of myo-inositol phosphate intermediates (Fig. 6.1). In addition, an acid phosphatase with phytate-degrading activity was identified in members of the Enterobacteriaceae family, such as E. coli (Cottrill et al., 2002), Pantoea agglomerans (Greiner, 2004) and E. cloacae (R. Greiner, 2005 unpublished data), which preferably degrades glucose1-phosphate. These enzymes were shown to hydrolyse only the phosphate residue at the D-3 position of phytate, pro-

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ducing D-myo-inositol 1,2,4,5,6-pentakisphosphate as the sole hydrolysis product. The alkaline phytate-degrading enzymes from cattail (Typha spp.; Hara et al., 1985), lily pollen (Lilium longiflorum; Barrientos et al., 1994) and Bacillus subtilis (Kerovuo et al., 2000) yield myo-inositol trisphosphate as the final product of phytate dephosphorylation (Fig. 6.2). With the exception of the phytate-degrading enzyme from L. longiflorum, alkaline phytate-degrading enzymes represent the class of β-propeller phosphatases. These seem to prefer the hydrolysis of every second phosphate rather than of adjacent ones, generating myo-inositol 2,4,6-trisphosphate and myo-inositol 1,3,5trisphosphate as the final dephosphorylation products (Kerovuo et al., 2000). Recent studies on the phytate-degrading enzymes of B. subtilis 168, B. amyloliquefaciens ATCC 15841 and B. amyloliquefaciens 45, however, point to myo-inositol 2,4,6trisphosphate as the sole final product of phytate degradation (R. Greiner, 2005 unpublished data). The alkaline phytate-degrading enzyme from L. longiflorum possesses the conserved active site motifs characteristic of histidine acid phosphatases (Mehta and Murthy, 2005) and generates a myo-inositol trisphosphate as the final phytate dephosphorylation product in a single degradation pathway by preferring removal of adjacent phosphate groups (Barrientos et al., 1994).

Regulation of phytate-degrading enzyme formation in microorganisms and plants In plant seeds and microorganisms, constitutive as well as inducible phytate-degrading enzymes have been identified. In microorganisms, expression of the inducible phytate-degrading enzymes is subjected to a complex regulation, but their formation is not controlled uniformly among different microorganisms (Konietzny and Greiner, 2004). Until now, phytate-degrading enzyme production was studied in some detail only in E. coli (Greiner et al., 1993), Raoultella terrigena (Greiner et al., 1997; Zamudio et al., 2002) and Saccharomyces cerevisiae (Andlid et al., 2004). A large increase in phytate-degrading activity was reported in germinating plant seeds and pollen, but the biochemical mechanisms leading to this rise in enzyme activity is still not well understood.

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I(1,2,3,4,5,6)P6 Barley P1, barley P2, spelt D12, wheat PHY1, wheat PHY2, wheat F2, oat, rice, rye, faba bean, lupine LP2, mung bean

D-I(1,2,3,5,6)P5

Wheat F2, mung bean(1)

Pseudomonas, Saccharomyces cerevisiae, Aspergillus ficuum, lupine LP11, lupine LP12, Pantoea agglomerans (2), Escherichia coli agp (2), Eenterobacter cloacae (2) D-I(1,2,4,5,6)P5

Barley P1, barley P2, spelt D12, wheat PHY1, wheat PHY2, oat, rice, rye, faba bean, lupine LP2

D-I(1,2,3,6)P4

I(1,2,3)P3

D-I(1,2,3,4,5)P5

E. coli appA

D-I(1,2,5,6)P4

D-I(1,2,6)P3

D-I(1,2)P2

E. coli appA, Paramecium

Paramecium

D-I(2,3,4,5)P4

D-I(1,2,3,4)P4

D-I(2,4,5)P3

I(1,2,3)P3

I(2,5)P2

D-I(2,3)P2

I(2)P Fig. 6.1. Major phytate degradation pathways by acid phytate-degrading enzymes barley P1, barley P2, spelt D12, rye, oat (Greiner and Larsson Alminger, 2001); wheat PHY1, wheat PHY2 (Nakano et al., 2000); rice (Hayakawa et al., 1990); faba bean, lupine LP11, lupine LP12, lupine LP2 (Greiner et al., 2002); wheat F2 (Lim and Tate, 1973); mung bean (Maiti et al., 1974); Saccharomyces cerevisiae (Greiner et al., 2001); Pseudomonas (Cosgrove, 1970); Escherichia coli (Greiner et al., 2000a; Cottrill et al., 2002); Paramecium (van der Kaay and van Haastert, 1995); Aspergillus ficuum (Chen and Li, 2003); Pantoea agglomerans (Greiner, 2004); Enterobacter cloacae (R. Greiner, 2005 unpublished data). (1) Generates also D-I(1,2,6)P3 and D-I(1,2)P2 as intermediates (2) D-I(1,2,4,5,6)P5 is the final product of phytate dephosphorylation

Synthesis in Microorganisms and Plants

81

I(1,2,3,4,5,6)P6 Lily

Bacillus subtilis

B. subtilis 168, B. amyloliquefaciens ATCC 15841, B. amyloliquefaciens 45

D-I(1,2,3,4,6)P5

D/L-I(1,2,3,4,5)P5

D/L-I(1,2,4,5,6)P5

D/L-I(1,2,3,4)P4

I(1,2,3,5)P4

I(2,4,5,6)P4

I(1,2,3)P3

I(1,3,5)P3

I(2,4,6)P3

D-I(1,2,4,5,6)P5

Fig. 6.2. Major phytate degradation pathways by the alkaline phytate-degrading enzymes Bacillus subtilis (Kerovuo et al., 2000); B. subtilis 168, B. amyloliquefaciens ATCC 15841, B. amyloliquefaciens 45 (from R. Greiner, 2005 unpublished data); lily (from Barrientos et al., 1994). The final phytate degradation product of cattail phytase was identified as myo-inositol trisphosphate (Hara et al., 1985), but no information was provided about the configuration of the generated phytate degradation products.

Regulation of phytate-degrading enzyme formation in microorganisms In non-limiting media, formation of the majority of the bacterial phytate-degrading enzymes was turned off in exponentially growing cells and started as soon as the cultures entered the stationary phase (Shimizu, 1992; Greiner et al., 1993, 1997; Sreeramulu et al., 1996; Choi et al., 1999; Zamudio et al., 2001; De Angelis et al., 2003). In moulds, phytate-degrading enzyme formation was growth-associated (Vats and Banerjee, 2002). Enzyme activity started to increase from the beginning of growth and continued to increase up to the onset of the stationary phase. As synthesis of bacterial phytate-degrading enzymes started as soon as the growth rate began to fall, it was suggested that either a nutrient or an energy limitation, both known to occur in the stationary phase, could cause their induction. Among the nutrient limitations tested, only carbon starvation was able to provoke an immediate synthesis of a phytate-degrading enzyme in R. terrigena (Greiner

et al., 1997). However, phytate-degrading enzyme formation in E. coli was triggered by phosphate starvation, while carbon, nitrogen and sulphur limitation were ineffective (Touati et al., 1987). A tight regulatory inhibition of the formation of phytatedegrading enzymes by phosphate levels was generally observed in all microorganisms, including moulds, yeast and bacteria, with the exception of R. terrigena (Greiner et al., 1997) and the rumen bacteria (Yanke et al., 1998), but only rarely could microorganisms utilize phytate as the sole source of carbon and phosphate. However, a small amount of phosphate in the growth medium probably stimulates phytate-degrading enzyme formation by enhancing microbial growth. Phosphate was shown to exert its effect on the synthesis of phytatedegrading enzymes at the level of transcription. The repression by phosphate seems to be less significant with more complex media (Greiner and Farouk, 2005), although it is not known which specific media components could account for this. Expression of phytate-degrading enzymes also depends on the nature of the carbon source,

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the initial pH and the temperature used for cultivation of the microorganisms. Both biomass and enzyme production respond to all these parameters, but temperature and pH for maximal growth and production of phytate-degrading enzymes were shown to be different in some moulds and yeast (Lambrechts et al., 1993; Kim et al., 1999; Sano et al., 1999; Mandviwala and Khire, 2000; Andlid et al., 2004), indicating a biomass-independent effect of temperature and pH on the synthesis of phytate-degrading enzymes in these microorganisms. No comparable studies have been performed so far in bacteria. In the presence of simple, easily fermentable sugars, strong repression of the formation of phytatedegrading enzymes was observed in several microorganisms, including moulds, yeast and bacteria (Shieh and Ware, 1968; Han and Gallagher, 1987; Lambrechts et al., 1993; Sano et al., 1999; Lan et al., 2002; Vats and Banerjee, 2002; Vohra and Satyanarayana, 2002). As with phosphate, an optimal concentration of glucose is also required. Low glucose levels result in low phytate-degrading activity due to reduced biomass production, whereas high levels inhibit enzyme production. However, the presence of glucose causes high levels of phytate-degrading activity in E. coli (Touati et al., 1987) and Lactobacillus amylovorus (Sreeramulu et al., 1996). The regulation of the formation of phytate-degrading enzymes was not suggested to be due to the carbon source itself, but to a change in the level of cellular cyclic adenosine monophosphate (cAMP). It has been established that a complex of cAMP with a protein called cAMP receptor protein (CRP) plays a central role in activating and repressing the expression of many genes (Saier et al., 1996). Cellular cAMP levels depend upon cell physiology, including the carbon source in the growth medium. That cAMP–CRP is directly involved in the regulation of the formation of phytate-degrading enzymes was shown in E. coli (Touati et al., 1987) and R. terrigena (Zamudio et al., 2002). In R. terrigena, synthesis of the phytate-degrading enzyme was downregulated by cAMP–CRP in the stationary phase and upregulated during exponential growth. Synthesis of phytate-degrading enzymes in E. coli has been reported to be downregulated by cAMP–CRP under all growth conditions. In moulds such as Aspergillus niger, formation of mycelial pellets in the presence of glucose or fructose as the sole carbon source was shown to be responsible for the low

enzyme yields (Shieh and Ware, 1968; Han and Gallagher, 1987). Dispersed growth and therefore an increase in phytate-degrading enzyme production could be obtained by using a medium containing a surfactant. For several Klebsiella spp. it was reported that phytate is needed to induce phytate-degrading enzyme production (Shah and Parekh, 1990; Tambe et al., 1994; Greiner et al., 1997), but the bacteria were unable to grow on phytate as the sole carbon source. Substrate induction was also found for Mitsuokella jalaludinii (Lan et al., 2002) and several Bacillus spp. (Powar and Jagannathan, 1982; Kerovuo et al., 1998; Kim et al., 1998), whereas phytate had no effect on the formation of phytate-degrading enzymes in E. coli (Greiner et al., 1993), Arxula adeninivorans (Sano et al., 1999) and Selenomonas ruminantium (Yanke et al., 1998). A reduced synthesis of the phytate-degrading enzymes in the presence of phytate was observed even in Schwanniomyces castellii (Lambrechts et al., 1993). An increased phosphate concentration in the growth medium due to phytate hydrolysis by the phytate-degrading enzymes secreted by the yeast could be responsible for this phenomenon. This suggestion would be in agreement with the observation that wheat and rice bran are excellent substrates for the production of extracellular phytate-degrading enzymes in microorganisms. As phytate in bran is less soluble than sodium phytate, phosphate concentrations are lower due to a slower release from bran phytate, and therefore repression of enzyme synthesis by phosphate is reduced. This ensures a continuous production of phytate-degrading enzymes during the whole fermentation process. The formation of phytate-degrading enzymes in Pseudomonas spp. (Irving and Cosgrove, 1971) and Klebsiella aerogenes (Tambe et al., 1994) was reported to be significantly induced in the presence of myo-inositol as the sole carbon source, although this was less effective than phytate. In the other Klebsiella spp. studied myo-inositol was ineffective (Shah and Parekh, 1990; Greiner et al., 1997). Anaerobiosis was effective in inducing phytatedegrading enzyme formation in E. coli (Greiner et al., 1993) and S. castellii (Lambrechts et al., 1993), whereas aeration had a positive effect on the production of phytate-degrading enzymes in A. ficcum (Nair et al., 1991). The presence of calcium ions in the growth medium was found to result in higher extracellular

Synthesis in Microorganisms and Plants

phytate-degrading activity in Bacillus spp. (Shimizu, 1992; Choi et al., 1999). However, the metal ions are not supposed to induce enzyme expression, but to stabilize the secreted enzyme. Binding of two calcium ions to high-affinity calcium binding sites was shown to result in a dramatic increase in thermostability of the phytate-degrading enzymes from Bacillus by joining loop segments remote in the amino acid sequence. Binding of three additional calcium ions to lowaffinity calcium binding sites at the top of the enzyme molecule turns on the catalytic activity of the enzyme by converting the highly negatively charged cleft into a favourable environment for the binding of phytate (Shin et al., 2001). Regulation of phytate-degrading enzyme formation on a molecular level was studied in detail in E. coli and S. cerevisae only.

Regulation of phytate-degrading enzyme formation in Escherichia coli At least two different phosphatases located in the periplasma of E. coli are capable of accepting myoinositol hexakisphosphate as a substrate. The agp-encoded acid phosphatase (EC 3.1.3.10) hydrolyses only the D-3 phosphate residue from phytate to produce D-myo-inositol 1,2,4,5,6-pentakisphosphate as the sole hydrolysis product (Cottrill et al., 2002). This enzyme has broad substrate specificity for phosphorylated compounds but demonstrates its highest activity towards glucose 1-phosphate, and it is believed that its function is to scavenge glucose (Pradel and Boquet, 1991). It appears to be largely synthesized constitutively, although small effects have been noted in regard to the amounts of this enzyme made under various growth conditions (Wanner, 1996). The appA-encoded phosphatase (EC 3.1.3.26) is highly specific for phytate and sequentially removes five of the six phosphate groups, starting with that attached to the D-6 position of the myo-inositol ring (Greiner et al., 2000a). Accumulation of this E. coli enzyme is controlled at the level of transcription and its expression is regulated by a complex regulatory mechanism involving several factors (Fig. 6.3). The appCBA operon contains the genes appA as well as cbdA and cbdB, coding for a putative cytochrome oxidase. These three genes are co-

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transcribed from a promoter (appC) located immediately upstream of cbdA and the operon, in addition, contains an internal promoter (appA) positioned upstream of appA (Atlung and Brøndsted, 1994). Both promoters are induced by phosphate starvation and by entry into the stationary phase (Atlung and Brøndsted, 1994), but the induction of the appC promoter is much stronger than that of the appA promoter. The induction by phosphate starvation has been shown to be independent of the PHO regulon (Touati et al., 1987), which is involved in the scavenging and specific uptake of phosphate from extracellular sources. No information on the regulation of the appA promoter under phosphate starvation and upon entry into stationary phase is available. In addition, the appC promoter was found to be activated by anaerobic growth conditions (Atlung and Brøndsted, 1994) and carbon starvation (Atlung et al., 1997). At low glucose levels the cAMP–CRP complex was suggested to directly interact with the appA promoter (Touati et al., 1987). Transcription from the appC promoter is dependent on the σS subunit of the RNA polymerase under all growth conditions tested, specifically during exponential growth, entry into the stationary phase in rich medium, starvation for carbon and phosphate and upon osmotic upshift (Atlung et al., 1997). It was suggested that σS affects expression of the appCBA operon directly. That σS controls the expression of genes responding to starvation, and cellular stress is well established. Therefore, the intracellular concentration of σS present in E. coli is influenced strongly by environmental factors. It increases upon entry into the stationary phase in rich medium, during starvation for carbon, nitrogen and phosphate (Gentry et al., 1993; Lange and Hengge-Aronis, 1994), and is increased strongly by osmotic upshift (Muffler et al., 1996). Although σS always affects the level of expression from the appC promoter, it is considered to have a regulatory role only during induction by osmotic upshift and upon entry into the stationary phase in rich medium, but the phosphate starvation-induced increase in σS concentration is not involved in the regulation of this operon. The expression of σS-dependent genes does not depend solely on the concentration of σS in the cell, but additional regulatory factors differentially modulate the expression of these genes.

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Phytase mRNA

PappC

cbdA

PappA

cbdB

Weak induction Strong induction

appA

mRNA

Ac t

iva

tio

n

AppY

AppY

PappY

X Induction

appY Activation by phosphorylation

ArcA P

ArcB cB Cell membrane

Fig. 6.3. Regulation of phytate-degrading enzyme formation in Escherichia coli. PappC = appC promoter; PappA = appA promoter; PappY = appY promoter; cbdA, cbdB = genes coding for a putative cytochrome oxidase; appA = gene encoding a phytate-degrading enzyme; appY = gene encoding AppY; AppY, ArcA = transcriptional regulators; ArcB = oxygen sensor; X = formate; P = phosphate.

Many σS-dependent genes are also under positive or negative control of other well-known global regulators such as cAMP–CRP, ArcA, AppY and Fnr. The appCBA operon was shown to be a target for the transcriptional activator AppY (Brøndsted and Atlung, 1996). The induction of the appC promoter by anaerobiosis is fully dependent on AppY, whereas its induction by phosphate starvation and upon entry into stationary phase (Fig. 6.3) is dependent strongly, but not exclusively, on this transcriptional regulator. AppY has no effect on the induction of the appA promoter. AppY expression itself is induced by anaerobiosis, starvation for phosphate and carbon, and upon entry into the stationary phase (Fig. 6.3), indicating that the increased AppY concentration under these conditions contributes to the increased expression of this E. coli phosphatase. An increase in σS was shown to be instrumental in the induction of appY during carbon starvation (Fig. 6.3).

The stationary phase induction of appY is only partially dependent on σS, whereas the induction during phosphate starvation and the growth rate regulation is independent of σS (Fig. 6.3). AppY-dependent induction of the appCBA operon during anaerobiosis and phosphate starvation seems to be mediated by a combination of increased appY expression and an activating signal for AppY generated under both conditions (Fig. 6.3). Formate, an intermediate in the mixed acid fermentation, was suggested to be this activating signal (Brøndsted and Atlung, 1996). However, the appC promoter also seems to be stimulated quite efficiently by AppY under non-activating conditions. In addition, appY expression always increased when glycerol was used as the carbon source. This was not due to a positive regulation by cAMP–CRP, but due to an increase in the doubling time in the presence of glycerol compared to glucose as a carbon source. That expression of appY is inversely correlated

Synthesis in Microorganisms and Plants

with the growth rate was shown previously (Brøndsted and Atlung, 1996). The ArcA response regulator, a second transcriptional regulator that is activated by the ArcB sensor in response to reduced respiration, activates transcription of the appCBA operon during entry into stationary phase and under anaerobic growth conditions (Fig. 6.3; Brøndsted and Atlung, 1996). During stationary phase induction, much of the ArcA effect is by AppY. It is possible that only the weak and early induction of the appCBA operon is mediated by ArcA in an AppY-independent manner. The signal that leads to the induction of the appC promoter upon entry into stationary phase may be primarily due to oxygen deprivation caused by an increase in cell density. AppY and ArcA depend on each other when activating the transcription of the appCBA operon during anaerobic growth. AppY expression was induced immediately by anaerobiosis, a process that is dependent on ArcA. The expression of the appCBA operon does not respond immediately to anaerobiosis but is delayed one generation, possibly due to the lack of sufficient AppY at the onset of anaerobiosis. Electron acceptors, which can be used in anaerobic respiration, repress the expression of the appCBA operon (Brøndsted and Atlung, 1996). The repression is particularly pronounced in the presence of nitrate. Since appY expression was not affected significantly by anaerobic energy metabolism (i.e. fermentation vs. anaerobic respiration), induction of the appCBA operon cannot be mediated by changes in the expression of AppY. It was suggested that nitrate repression was partially dependent on NarL, which activates transcription of operons involved in nitrate respiration and represses the synthesis of alternate respiratory enzymes (Berg and Stewart, 1990). The residual nitrate repression could be mediated by NarP (Rabin and Stewart, 1993). Alternatively, the residual anaerobic repression by nitrate and the repression by fumarate could be indirect effects of ArcA, as the level of active ArcA is dependent on the respiratory state of the cell. Regulation of phytate-degrading enzyme formation in Saccharomyces cerevisiae In the yeast S. cerevisiae, the PHO regulon controls expression of the PHO genes at the transcription

85

level depending on the extracellular phosphate concentration (Lemire et al., 1985), but it is not known how extracellular phosphate levels are detected by the yeast. The PHO gene family is involved in the scavenging and specific uptake of phosphate from extracellular sources and at least three phosphatases, encoded by PHO5, PHO10 and PHO11, are capable of hydrolysing myo-inositol hexakisphosphate (Andlid et al., 2004). All these enzymes are extracellular oligomeric glycoproteins with an acidic pH optimum and broad substrate specificity. PHO5 encodes the major contributor to the secreted acid phosphatase activity, whereas PHO10 and PHO11 encode only a minor fraction (Lemire et al., 1985). The information about the extracellular phosphate level is transmitted to the phosphatase encoding genes by a set of positive and negative regulatory proteins, which are encoded by at least the following genes: PHO2, PHO4, PHO80, PHO81 and PHO85. Besides PHO5, PHO10 and PHO11, at least the following additional genes are known to be regulated by the PHO regulon: PHO8 encoding a non-specific repressible alkaline phosphatase localized in the vacuole; PHO84 and PHO89 encoding high-affinity phosphate transporters localized in the plasma membrane; GIT1 encoding a glycerophosphoinositol transporter localized in the plasma membrane; PHO86 encoding a protein required for the correct localization of Pho84p in the plasma membrane; PHO81, SPL2 and YPL110C encoding regulatory proteins, and several genes encoding various phosphate metabolism enzymes such as the PHM genes that are involved in polyphosphate synthesis or degradation (Pinson et al., 2004). For the transcriptional activation of PHO5, PHO10 and PHO11 the two DNA-binding proteins Pho4p, encoded by PHO4, and Pho2p, encoded by PHO2, are needed (Fig. 6.4). Pho4p binds to a specific cis-acting regulatory site (UAS) in the promoter of all PHO genes (Oshima, 1997), and Pho2p forms a ternary complex with Pho4p on a PHO promoter (Barbari´c et al., 1996). However, Pho2p does not have a direct function in the transduction of the extracellular phosphate levels, but interaction of the Pho2p/Pho4p/UAS ternary complex with basal transcription factors is considered to initiate transcription of the PHO genes (Magbanua et al., 1997). Both PHO4 and PHO2 are transcribed at low levels (Yoshida et al., 1989).

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PPHO Pho4p

PHO-regulated gene

Pho2p

mRNA

Nuclear membrane Pho2p

Pho81p inhibitor

Pho80p cyclase Pho85p kinase

Phytase

Pho4p Pho84p Pho89p Pho89p

Cell membrane Pho84p

Fig. 6.4. Regulation of phytate-degrading enzyme formation in Saccharomyces. PPHO = PHO promoter; Pho2p, Pho4p = regulatory proteins (DNA-binding proteins); Pho81p = regulatory protein (inhibitor); Pho80p = cyclase; Pho85p = kinase; Pho84p, Pho89p = high-affinity phosphate transporters; PHO regulated genes = PHO5, PHO10 and PHO11 encoding repressible non-specific acid phosphatases; PHO8 encoding a non-specific repressible alkaline phosphatase; PHO84 and PHO89 encoding highaffinity phosphate transporters; GIT1 encoding a glycerophosphoinositol transporter; PHO86 encoding a protein required for the correct localization of Pho84p; PHO81, SPL2 and YPL110C encoding regulatory proteins; several genes encoding various phosphate metabolism enzymes such as the PHM genes.

PHO4 transcription is constitutive, whereas PHO2 transcription is self-regulated. High extracellular phosphate concentrations result in phosphorylation of five serine residues in Pho4p by a complex of the negative regulators Pho80p and Pho85p (Kaffman et al., 1994). This phosphorylated Pho4p is not imported into the nucleus (O’Neill et al., 1996) and therefore transcription of PHO genes is turned off (Fig. 6.4). Pho81p, which is inactive under high extracellular phosphate concentrations, is activated when the extracellular phosphate level is sufficiently low. Activated Pho81p inhibits the function of the Pho80p–Pho85p complex (Ogawa et al., 1995), thus allowing translocation of Pho4p into the nucleus, where Pho4p, together with Pho2p, activates the transcription of the PHO genes (Fig. 6.4; Komeili and O’Shea, 1999). PHO80 and PHO85 are transcribed constitutively at low levels (Madden et al., 1990), whereas transcription

of PHO81 is regulated by the level of extracellular phosphate by the same PHO regulon, indicating that the regulatory circuit forms a positive feedback loop (Ogawa et al., 1993).

Regulation of phytate-degrading enzyme formation in plants Studies in plant seeds and pollen indicate that there are constitutive and germination-inducible phytate-degrading enzymes. Thus, two main mechanisms appear to be involved in the regulation of phytate breakdown during germination: control of the activity of the hydrolytic enzymes and control of their rate of synthesis. Constitutive phytate-degrading enzymes, which are found in all seeds and pollen, are present in a fully active form in mature seeds and pollen, and are consid-

Synthesis in Microorganisms and Plants

ered to start phytate breakdown during imbibition. Germination-inducible phytate-degrading enzymes are synthesized de novo either from a long-lived, pre-existing mRNA as in lily (Lin et al., 1987) and petunia (Jackson and Linskens, 1982), or by regulating enzyme synthesis at the level of transcription as in wheat (Bianchetti and Sartirana, 1967), barley (Katayama and Suzuki, 1980), maize (Maugenest et al., 1999), pea (Kuvaeva and Kretovich, 1978), mung bean (Mandal and Biswas, 1970) and lentils (Greiner et al., 2005). In addition, there is good evidence for the activation of pre-existing inactive phytatedegrading enzymes during early stages of germination (Eastwood and Laidman, 1971; Gabard and Jones, 1986). Many alternative ways can be suggested by which enzymes can be reversibly inactivated: folding; enclosure in, or attachment to, membranes; association or dissociation of subunits; or addition of a section to the polypeptide chain of the active protein. In such cases, during germination a reversal of these processes could occur. Gibberellic acid and phosphate may control phytate degradation during germination, but so far there is only a small amount of contradictory information available on the effect of both compounds on phytate-degrading activity in plant seeds and pollen. In maize, gibberellic acid does not significantly alter the accumulation of phytate-degrading enzymes during germination (Maugenest et al., 1999), but no information about its effect on phytate breakdown is available. An enhancement of phytate degradation by gibberellic acid without any effect on measurable phytate-degrading activity was found in barley (Gabard and Jones, 1986) and wheat (Eastwood and Laidman, 1971). Gabard and Jones (1986) suggested that gibberellic acid merely increases the secretion of phytate-degrading enzymes, but does not stimulate their synthesis, thus giving phytate-degrading enzymes access to phytate. Eastwood and Laidman (1971) claimed that gibberellic acid stimulates phytate breakdown by releasing phosphate, a potent competitive inhibitor of many phytate-degrading enzymes (Konietzny and Greiner, 2002), from aleurone cells, where most of the cereal phytate-degrading activity is located. In barley (Katayama and Suzuki, 1980) and lentils (Greiner et al., 2005), gibberellic acid was shown to enhance phytatedegrading activity and phytate degradation dur-

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ing germination and it was concluded that this effect of gibberellic acid is due at least in part to a stimulation of de novo enzyme synthesis participating in phytate breakdown. Centeno et al. (2001) reported an increase in phytate-degrading activity accompanied by a reduction in phytate breakdown in rye and barley in the presence of gibberellic acid, but gave no interpretation for this apparently contradictory observation. Two main mechanisms appear to be involved in the regulation of phytate-degrading activity by phosphate. Most phytate-degrading enzymes are strongly inhibited by phosphate in vitro (Konietzny and Greiner, 2002), so enzyme activity itself may also be controlled by phosphate in vivo (Eastwood and Laidman, 1971; Katayama and Suzuki, 1980). Furthermore, it was concluded that phosphate also acts at the transcription level through repression of phytate-degrading enzyme expression (Sartirana and Bianchetti, 1967). One explanation for the inconsistency of the available data on the regulation of phytatedegrading activity in plant seeds and pollen during germination is the presence of several molecular forms of phytate-degrading enzymes in a certain plant (Kuvaeva and Kretovich, 1978; Goel and Sharma, 1979; Baldi et al., 1988; Konietzny et al., 1995; Hamada, 1996; Maugenest et al., 1999; Greiner et al., 2000b; Greiner, 2002) that are regulated in different ways and may have different physiological functions. Different analytical approaches to determine phytate-degrading activity may also contribute to the conflicting findings. It was previously shown that values obtained by extraction methods are considerably lower than those obtained by direct incubation methods (Greiner and Egli, 2003).

In vivo Function of Phytatedegrading Enzymes Phytate-degrading enzymes in plant seeds and microorganisms occur in multiple forms (Kuvaeva and Kretovich, 1978; Goel and Sharma, 1979; Baldi et al., 1988; Hamada, 1994; Konietzny et al., 1995; Maugenest et al., 1999; Greiner et al., 2000b; Cottrill et al., 2002; Greiner, 2002). These forms may exhibit different stereospecificity of phytate dephosphorylation, be regulated in different ways, be directed to different localization

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within and outside the producing cell, and thus may have different physiological functions. As the classification of ‘phytases’ is solely according to their capability of hydrolysing phytate in vitro, some of these enzymes may not be involved in phytate degradation in vivo but may have completely different functions.

Proposed functions of phytate-degrading enzymes in plants Phytate-degrading enzymes in higher plants occur predominantly in grains, seeds and pollen, but little information is available on their specific localization within these. In cereal seeds, phytate-degrading activity was found to be mainly associated with the aleurone layer (Gabard and Jones, 1986), whereas in legume seeds it was reported to be located in the cotyledons (Gibson and Ullah, 1988; Hegeman and Grabau, 2001). Further, the extraction of phytate-degrading activity is strongly enhanced by the presence of Triton X-100, suggesting an association with membrane structures (Scott and Loewus, 1986; Scott, 1991; Greiner and Egli, 2003). The germination-inducible enzymes in particular are responsible for phytate breakdown during germination to make phosphate, minerals and myo-inositol available for plant growth and development (Greiner et al., 2005). Some phytate-degrading activity is also found in plant roots (Hübel and Beck, 1996; Li et al., 1997; Hayes et al., 1999), in which it may play a role in providing the central stele with minerals (Maugenest et al., 1999) or allow plants to use soil inositol phosphates. However, phytate appears to be only poorly utilized by plants, probably due to a combination of low phytatedegrading activity of roots and the low solubility of soil phytate (Hayes et al., 2000; see also Richardson et al., Chapter 15, this volume). Thus, it was suggested that soil microorganisms colonizing the plant rhizosphere and producing extracellular phytate-degrading activity, such as Bacillus and Enterobacter ssp., or membrane-bound phytate-degrading enzymes such as those synthesized by mycorrhizal fungi, could act as plant growth promoting microorganisms by making phytate phosphate available to the plant (McElhinney and Mitchell, 1993; Richardson et al., 2001; Idriss et al., 2002).

Possible roles of phytate-degrading enzymes in microbes The efficient de-repression of phytate-degrading enzyme formation by phosphate starvation in most microorganisms suggests a possible role for these enzymes in providing the cell with phosphate. This is supported by the identification of a phytate-degrading enzyme in the stalk of Caulobacter crescentus, an aquatic bacterium that lives in oligotrophic environments where phosphate limits productivity (Ireland et al., 2002). Phosphate uptake is one of the hypothesized functions of the stalks, which is enhanced when the stalks elongate during phosphate limitation. This increase in surface area, as well as the presence of a phytate-degrading enzyme, would allow the uptake of organic phosphate. With respect to phytate utilization it is also worth mentioning that cyanobacteria belonging to Rivulariaceae showed better growth in phytate when they form hairs compared to non-hair-forming Rivulariaceae (Whitton et al., 1991). The assumption could also explain why, with the exception of sourdough bacteria, there is no clear evidence for lactic acid bacteria with the ability to degrade phytate. Lactic acid bacteria are adapted to environments rich in nutrients and energy where evolutionary selection pressure would not favour the capability to produce a phytate-degrading enzyme. Phytate-degrading enzymes in S. cerevisiae are part of the PHO protein family, which is involved in the scavenging and specific uptake of phosphate from extracellular sources (Lemire et al., 1985). In E. coli, however, the phytate-degrading enzymes are not under the control of the PHO regulon (Touati et al., 1987). Therefore, they do not appear to have a primary role in phosphate assimilation, even though they probably contribute to periplasmatic phosphate levels under certain conditions. The agp-encoded phosphatase acts primarily as a glucose scavenger, whereas the appA-encoded phosphatase is believed to have a role in stress protection. In E. coli, the primary response to the limitation of a specific nutrient is activation of a certain set of genes that improves uptake of the nutrient present in low concentration or allows the utilization of other substances that belong to the same class of nutrient. These nutrient-specific systems include the cAMP–CRP regulon for the use of alternative carbon sources, the NtrB/NtrC/σ54 regulon that is induced

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under nitrogen limitation and the PhoB/PhoR regulon that is induced under phosphorus limitation, as well as systems for scavenging low concentrations of iron and other essential substances (Hengge-Aronis, 1996). If alternative nutrient sources are present in the growth medium, the cells continue to grow and divide. However, if the environment is totally exhausted of an essential nutrient, the cells enter into the stationary phase. In contrast to the specific responses, the nature of the stationary phase response does not appear to be dependent on the limiting nutrient. Thus, the reaction to nutrient limitation can be seen as a two-stage process. If the induction of the nutrient-specific responses remains unsuccessful (i.e. growth cannot be resumed), the stationary phase response is included. This secondary response involves a transition from a metabolic state aimed at maximal growth and cell division to a maintenance metabolism and the induction of many genes whose function it is to provide maximal protection against a large variety of stress conditions. In contrast to most other microorganisms, anaerobic rumen bacteria are capable of tolerating a high level of phosphate without any negative impact on phytate-degrading enzyme formation (Yanke et al., 1998). This unique ability may result in efficient phytate hydrolysis in the rumen, even under the high phosphate levels in the rumen fluid of ruminants fed concentrated feed. However, this raises the question about the physiological function of this bacterial phytate-degrading activity if phosphate limitation is not a problem.

Alternative functions of phytatedegrading enzymes To provide the cell with phosphate, phytatedegrading enzymes must have access to phytate in the environment. Extracellular phytatedegrading enzymes have been identified in moulds and yeast, whereas in bacteria these enzymes are mainly cell-associated. The only bacteria showing extracellular phytate-degrading activity are those of the genera Bacillus (Powar and Jagannathan, 1982; Shimizu, 1992; Kerovuo et al., 1998; Kim et al., 1998) and Enterobacter (Yoon et al., 1996). The phytate-degrading enzymes of E. coli have been reported to be

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periplasmatic proteins (Greiner et al., 1993). However, phytate is equally well hydrolysed by both disrupted and intact E. coli cells (R. Greiner, 2005 unpublished data). Consequently these enzymes appear to have free access in vivo to the substrates present in the surrounding medium. Phytate-degrading activity in S. ruminantium and M. multiacidus was found to be associated with the outer membrane, even though phosphate was released from phytate into the culture fluid by pure cultures (D’Silva et al., 2000). It is not known how bacteria with an apparent lack of extracellular phytate-degrading activity, such as some Pseudomonas strains, either grow in the absence of a readily utilizable phosphate source or acquire phosphate. As no phosphate was detected in the growth medium either initially or throughout the growth period (Richardson and Hadobas, 1997), phytate might be transported into the bacterial cells. Wang et al. (2004) suggested, for example, that in K. pneumoniae the gene encoding the phytate-degrading enzyme is co-transcribed from a polycistronic mRNA, which also acts as a template for an inositol phosphate transporter. The role of phytate-degrading enzymes is not limited to simple degradation functions in metabolic pathways. In fact, myo-inositol phosphate phosphatase activities were shown to be involved in signal transduction, cell division and microbial pathogenesis (Craxton et al., 1997; Zhou et al., 2001). The discovery of myo-inositol fluxes as a result of pathogen infections is an intriguing finding, particularly as phospholipid signalling plays a critical role in many host cell functions, including those affecting cellular survival and regulation of intracellular membrane trafficking (DeVinney et al., 2000). Although several pathogens trigger myo-inositol phosphate fluxes, different mechanisms appear to be involved. Many gram-negative animal and plant pathogens have developed specialized type III secretion systems to translocate bacterial proteins directly into the eukaroytic host cell (DeVinney et al., 2000). These can influence survival, internalization and replication of the pathogens. For enteropathogenic E. coli, Helicobacter pylori and Salmonella typhimurium it was proposed that intimate contact between the pathogen and the host cell stimulates host cell phospholipase C activity, resulting in the cleavage of phosphatidylinositol 4,5-bisphosphate into the second messengers

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1,4,5-trisphosphate and diacylglycerol (Ruschkowski et al., 1992; Foubister et al., 1994). For Listeria monocytogenes, a secreted phosphatidylinositol-specific phospholipase C (PI-PLC) was suggested to be an essential determinant of their pathogenesis (Camilli et al., 1991). A number of further gram-positive human pathogens secrete PI-PLCs, including Staphylococcus aureus, Clostridium novyi, B. cereus and B. anthracis (Low, 1989). Salmonella dublin was reported to modulate host cell signalling pathways by translocating at least two proteins, SopB and SopE (Zhou et al., 2001). SopB exhibits myoinositol phosphate phosphatase activity and specifically dephosphorylates D-myo-inositol 1,3,4,5,6-pentakisphosphate to D-myo-inositol 1,4,5,6-tetrakisphosphate. A phosphatase capable of catalysing the same reaction was also found in other Enterobacteriaceae family members, such as E. coli (Cottrill et al., 2002), P. agglomerans (Greiner, 2004) and E. cloacae (R. Greiner, 2004 unpublished data). Due to the unique myo-inositol phosphate phosphatase activity of these enzymes, Cottrill et al. (2002) raised the question of a role for them in microbial pathogenesis or cellular myo-inositol phosphate metabolism. However, in contrast to SopB (Norris et al., 1998), these three enzymes also hydrolyse phytate in vitro. SopE has no inherent phosphatase activity, but is proposed to activate an endogenous cellular myo-inositol phosphate phosphatase (Zhou et al., 2001). In addition, both SopB and SopE stimulate cellular responses that lead to host cell phospholipase C activation. Xanthomonas oryzae pv. oryzae, an important rice pathogen, has been suggested to require a Bacillus-like phytate-degrading enzyme for optimal virulence (Chatterjee et al., 2003). Homologues of this gene are also present in the genomes of X. campestris pv. campestris and X. axonopodis pv. citri. The enzyme is secreted and may interfere with myo-inositol phosphate-based signalling processes in plants. Differences between this Xanthomonas protein and the Bacillus phytate-degrading enzyme, specifically in residues involved in the binding of the phosphate residue attached to the D-2 position of the phytate molecule (Chatterjee et al., 2003), may result in an inability to bind phytate and other myoinositol phosphates phosphorylated at the D-2 position. Therefore, this Xanthomonas enzyme may exhibit a myo-inositol phosphate phosphatase

activity that emulates the virulence-associated SopB phosphatase from S. dublin. Position D-2 is the only one in axial orientation, and all myo-inositol phosphate intermediates identified so far during enzymatic phytate breakdown are phosphorylated in this position. However, with the exception of phytate itself, intracellular myo-inositol phosphates are always dephosphorylated at the D-2 position. The cell may therefore discriminate between intra- and extracellular-generated myo-inositol phosphates by the phosphorylation status of the D-2 position of the myo-inositol ring. Further, a considerable body of evidence supports the hypothesis that the phosphorylation status of the D-2 position is used to independently control the synthesis and breakdown of phytate in the plant kingdom. For example, this was concluded from the myo-inositol phosphates identified in a development stage of the plant Spirodela polyrhiza L., which is associated with massive accumulation of phytate (Brearly and Hanke, 1996a,b). None of these myo-inositol phosphates was phosphorylated at the D-2 position of the myo-inositol ring.

Conclusions and Future Research Directions Until now, research on phytate-degrading enzymes has focused almost exclusively on making enzymes available that are suitable for use as animal feed additives. The in vivo function of phytate-degrading enzymes has therefore received little attention. So far, only the germinationinducible phytate-degrading enzymes of plant seeds could be called phytases. Their action upon phytate is considered likely, because the breakdown of phytate during germination makes phosphate, minerals and myo-inositol available for plant growth and development. As formation of extracellular phytate-degrading enzymes in moulds and yeast is triggered by phosphate starvation, these enzymes hydrolyse organic phosphates, including phytate, to provide the cell with phosphate from extracellular sources. These enzymes are therefore non-specific phosphatases that exhibit phytate-degrading activity. The in vivo function of other phytate-degrading enzymes is mainly speculative. In addition to providing the cell with phosphate as mentioned

Synthesis in Microorganisms and Plants

above, a role in stress response or bacterial pathogenesis has been assumed. Information on the regulation of phytatedegrading enzyme formation might shed light on the in vivo role of phytate-degrading enzymes. Extensive studies have so far been performed only for E. coli and S. cerevisiae, although some information on the regulation of phytate-degrading enzyme formation is also available for R. terrigena and germinating cereal and legume seeds. The majority of studies, however, were performed on a trial-and-error basis to optimize medium composition with respect to phytatedegrading enzyme yields. Micro-arrays should be used to learn more about the regulation of phytate-degrading enzyme formation and their in vivo functions, in addition to classical biochemical and molecular approaches. Running transcriptome projects may provide useful information on the expression of genes encoding phytate-degrading enzymes. There are already large data-sets, e.g. for yeast (http://www.transcriptome.ens.fr/ymgv) and Arabidopsis (https://www.genevestigator.

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ethz.ch), online which could be mined for the information they contain on the environmental and biotic factors that contribute to expression of genes encoding phytate-degrading enzymes. In addition to understanding the regulation of phytate-degrading enzyme formation and their in vivo function, a number of other important questions remain to be answered. Only a relatively small number of organisms have been identified as being able to utilize phytate as the sole phosphate or phosphate and carbon source, but the prevalence of this ability among microbes remains unknown. More information is required on the ecology of phytate-utilizing bacteria, where they occur, and the site of phytate dephosphorylation. Furthermore, it has not been clearly established that only extracellular phytatedegrading enzymes make phytate-phosphate available to the microbial cell. It therefore remains possible that phytate is transported intact across the microbial cell wall to be dephosphorylated within the cell.

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Kim, Y.-O., Kim, H.-K., Bae, K.-S., Yu, J.-H. and Oh, T.-K. (1998) Purification and properties of a thermostable phytase from Bacillus sp. DS11. Enzyme and Microbial Technology 22, 2–7. Kim, D.-S., Godber, J.S. and Kim, H.-R. (1999) Culture conditions for a new phytase-producing fungus. Biotechnology Letters 21, 1077–1081. Komeili, A. and O’Shea, E.K. (1999) Roles of phosphorylation sites in regulating activity of the transcription factor Pho4. Science 284, 977–980. Konietzny, U. and Greiner, R. (2002) Molecular and catalytic properties of phytase-degrading enzymes (phytases). International Journal of Food Science and Technology 37, 791–812. Konietzny, U. and Greiner, R. (2004) Bacterial phytase: potential application, in vivo function and regulation of its synthesis. Brazilian Journal of Microbiology 35, 11–18. Konietzny, U., Greiner, R. and Jany, K.-D. (1995) Purification and characterization of a phytase from spelt. Journal of Food Biochemistry 18, 165–183. Kuvaeva, E.B. and Kretovich, V.L. (1978) Phytase of germinating pea seeds. Soviet Plant Physiology 25, 290–295. Lambrechts, C., Boze, H., Segueilha, L., Moulin, G. and Galzy, P. (1993) Influence of culture conditions on the biosynthesis of Schwanniomyces castellii phytase. Biotechnology Letters 15, 399–404. Lan, G.Q., Abdullah, N., Jalaludin, S. and Ho, Y.W. (2002) Culture conditions influencing phytase production of Mitsuokella jalaludinii, a new bacterial species from the rumen of cattle. Journal of Applied Microbiology 93, 668–674. Lange, R. and Hengge-Aronis, R. (1994). The cellular concentration of the σS subunit of the RNA polymerase in Escherichia coli is controlled at the level of transcription, translation, and protein stability. Genes and Development 8, 1600–1612. Lemire, J.M., Willcocks, T., Halvorson, H.O. and Bostian, K.A. (1985) Regulation of repressible acid phosphatase gene transcription in Saccharomyces cerevisiae. Molecular and Cellular Biology 5, 2131–2141. Li, M., Osaki, M., Honma, M. and Tadano, T. (1997) Purification and characterization of phytase induced in tomato roots under phosphorus-deficient conditions. Soil Science and Plant Nutrition 43, 179–190. Lim, P.E. and Tate, M.E. (1973) The phytases. II. Properties of phytase fraction F1 and F2 from wheat bran and the myo-inositol phosphates produced by fraction F2. Biochimica et Biophysica Acta 302, 326–328. Lin, J.-J., Dickinson, D.B. and Ho, T.-H.D. (1987) Phytic acid metabolism in lily (Lilium longiflorum Thunb.) pollen. Plant Physiology 83, 408–413. Low, M.G. (1989) The glycosyl-phosphatidylinositol anchor of membrane proteins. Biochimica et Biophysica Acta 988, 427–454. Madden, S.L., Johnson, D.L. and Bergman, L.W. (1990) Molecular and expression analysis of the negative regulators involved in the transcriptional regulation of acid phosphatase production in Saccharomyces cerevisiae. Molecular and Cellular Biology 10, 5950–5957. Magbanua, J.P.V., Ogawa, N., Harashima, S. and Oshima, Y. (1997) The transcriptional activators of the PHO regulon, Pho4p and Pho2p, interact directly with each other and with components of the basal transcription machinery in Saccharomyces cerevisae. Journal of Biochemistry 121, 1182–1189. Maiti, I.B., Majumber, A.L. and Biswas, B.B. (1974) Purification and mode of action of phytase from Phaseolus aureus. Phytochemistry 13, 1047–1051. Mandal, N.C. and Biswas, B.B. (1970) Metabolism of inositol phosphates. 1. Phytase synthesis during germination in cotyledons of mung beans, Phaseolus aureus. Plant Physiology 45, 4–7. Mandviwala, T.N. and Khire, J.M. (2000) Production of high activity thermostable phytase from thermotolerant Aspergillus niger in solid state fermentation. Journal of Industrial Microbiology and Biotechnology 24, 237–243. Maugenest, S., Martinez, I., Godin, B., Perez, P. and Lescure, A.-M. (1999) Structure of two maize phytase genes and their spatio-temporal expression during seedling development. Plant Molecular Biology 39, 502–514. McElhinney, C. and Mitchell, D.T. (1993) Phosphatase activity of four ectomycorrhizal fungi found in a Sitka spruce–Japanese larch plantation in Ireland. Mycological Research 97, 725–732. Mehta, B.D. and Murthy, P.P.N. (2005) Unique alkaline phytase from lily pollen: cloning, characterization and differential expresssion. In: Turner, B.L., Richardson, A.E. and Mullaney, E. J. (eds) Inositol Phosphates in the Soil–Plant–Animal System: Linking Agriculture and Environment. Proceedings of the Bouyoucos Conference on Inositol Phosphates in the Environment, 21–24 August 2005, Sun Valley, Idaho, pp. 70. Muffler, A., Traulsen, D.D., Lange, R. and Hengge-Aronis, R. (1996) Posttranscriptional osmotic regulation of the σS subunit of RNA polymerase in Escherichia coli. Journal of Bacteriology 178, 1607–1613. Nair, V.C., Laflamme, J. and Duvnjak, Z. (1991) Production of phytase by Aspergillus ficuum and reduction of phytic acid content in canola meal. Journal of the Science of Food and Agriculture 54, 355–365. Nakano, T., Joh, T., Narita, K. and Hayakawa, T. (2000) The pathway of dephosphorylation of myo-inositol hexakisphosphate by phytases from wheat bran of Triticum aestivum L. cv. nourin #61. Bioscience, Biotechnology and Biochemistry 64, 995–1003.

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Norris, F.A., Wilson, M.P., Wallis, T.S., Galyov, E.E. and Majerus, P.W. (1998) SopB, a protein required for virulence of Salmonella dublin, is an inositol phosphate phosphatase. Proceedings of the National Academy of Sciences of the United States of America 95, 14057–14059. Ogawa, N., Noguchi, K., Yamashita, Y., Yasuhara, T., Hayashi, N., Yoshida, K. and Oshima, Y. (1993) Promoter analysis of the PHO81 gene encoding a 134 kDa protein bearing ankyrin repeats in the phosphatase regulon of Saccharomyces cerevisiae. Molecular General Genetics 238, 444–454. Ogawa, N., Noguchi, K.-I., Sawai, H., Yamashita, Y., Yompakdee, C. and Oshima, Y. (1995) Functional domains of Pho81p, an inhibitor of the Pho85p protein kinase, in the transduction pathway for Pi signals in Saccharomyces cerevisiae. Molecular and Cellular Biology 15, 997–1004. O’Neill, E.M., Kaffman, A., Jolly, E.R. and O’Shea, E.K. (1996) Regulation of PHO4 nuclear localization by the PHO80–PHO85 cyclin–CDK complex. Science 271, 95–99. Oshima, Y. (1997) The phosphatase system in Saccharomyces cerevisiae. Genes and Genetic Systems 72, 323–334. Pinson, B., Merle, M., Franconi, J.-M. and Daignam-Fornier, B. (2004) Low affinity orthophosphate carriers regulate PHO gene expression independently of internal orthophosphate concentration in Saccharomyces cerevisiae. Journal of Biological Chemistry 279, 35273–35280. Powar, V.K. and Jagannathan, V. (1982) Purification and properties of phytate-specific phosphatase from Bacillus subtilis. Journal of Bacteriology 151, 1102–1108. Pradel, E. and Boquet, P.L. (1991) Utilization of exogenous glucose-1-phosphate as a source of carbon or phosphate by Escherichia coli K12: respective roles of acid glucose-1-phosphatase, hexosephosphate permease, phosphoglucomutase and alkaline phosphatase. Research in Microbiology 142, 37–45. Rabin, R.S. and Stewart, V. (1993) Dual response regulators (NarL and NarP) interact with dual sensors (NarX and NarQ) to control nitrate- and nitrite-regulated gene expression in Escherichia coli K-12. Journal of Bacteriology 175, 3259–3268. Richardson, A.E. and Hadobas, P.A. (1997) Soil isolates of Pseudomonas spp. that utilize inositol phosphates. Canadian Journal of Microbiology 43, 509–516. Richardson, A.E., Hadobas, P.A., Hayes, J.E., O’Hara, C.P. and Simpson, R.J. (2001) Utilization of phosphorus by pasture plants supplied with myo-inositol hexakisphosphate is enhanced by the presence of soil microorganisms. Plant and Soil 229, 47–56. Ruschkowski, S., Rosenshine, I. and Finlay, B.B. (1992) Salmonella typhimurium induces an inositol phosphate flux in infected epithelial cells. FEMS Microbiological Letters 74, 121–126. Saier, M.H. Jr, Ramseier, T.M. and Reizer, J. (1996) Regulation of carbon utilization. In: Neidhardt, F.C., Curtiss, R. III, Ingraham, J.L., Lin, E.C.C., Low, K.B., Magasanik, B., Reznikoff, W.S., Riley, M., Schaechter, M. and Umbarger, H.E. (eds) Escherichia coli and Salmonella: Cellular and Molecular Biology, 2nd edn. ASM Press, Washington, DC, pp. 1325–1343. Sano, K., Fukuhara, H. and Nakamura, Y. (1999) Phytase of the yeast Arxula adeninivorans. Biotechnology Letters 21, 33–38. Sartirana, M.L. and Bianchetti, R. (1967) The effect of phosphate on the development of phytase in the wheat embryo. Physiologia Plantarum 20, 1066–1075. Scott, J. J. (1991) Alkaline phytase activity in nonionic detergent extracts of legume seeds. Plant Physiology 95, 1298–1301. Scott, J. J. and Loewus, F.A. (1986) A calcium-activated phytase from pollen of Lilium longiflorum. Plant Physiology 82, 333–335. Shah, V. and Parekh, L. J. (1990) Phytase from Klebsiella sp. no. PG-2: purification and properties. Indian Journal of Biochemistry 27, 98–102. Shieh, T.R. and Ware, J.H. (1968) Survey of microorganisms for the production of extracellular phytase. Applied Microbiology 16, 1348–1351. Shimizu, M. (1992) Purification and characterization of a phytase from Bacillus subtilis (Natto) N-77. Bioscience, Biotechnology, and Biochemistry 56, 1266–1269. Shin, S., Ha, N.-C., Oh, B.-C., Oh, T.-K. and Oh, B.-H. (2001) Enzyme mechanism and catalytic property of β propeller phytase. Structure 9, 851–858. Sreeramulu, G., Srinivasa, D.S., Nand, K. and Joseph, R. (1996) Lactobacillus amylovorus as a phytase producer in submerged culture. Letters in Applied Microbiology 23, 385–388. Tambe, S.M., Kaklij, G.S., Keklar, S.M. and Parekh, L.J. (1994) Two distinct molecular forms of phytase from Klebsiella aerogenes: evidence for unusually small active enzyme peptide. Journal of Fermentation and Bioengineering 77, 23–27. Touati, E., Dassa, E., Dassa, J. and Boquet, P.L. (1987) Acid phosphatase (pH 2.5) of Escherichia coli: regulatory characteristics. In: Torriani-Gorini, A., Rothman, F.G., Silver, S., Wright, A. and Yagil, E. (eds) Phosphate Metabolism and Cellular Regulation in Microorganisms. American Society for Microbiology, Washington, DC, pp. 31–40.

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van der Kaay, J. and van Haastert, J.M. (1995) Stereospecificity of inositol hexaphosphate dephosphorylation by Paramecium phytase. Biochemical Journal 312, 907–910. Vats, P. and Banerjee, U.C. (2002) Studies on the production of phytase by a newly isolated strain of Aspergillus niger var tiegham obtained from rotten wood-logs. Process Biochemistry 38, 211–217. Vohra, A. and Satyanarayana, T. (2002) Statistical optimization of the media components by response surface methodology to enhance phytase production by Pichia anomala. Process Biochemistry 37, 999–1004. Wang, X., Upatham, S., Panbangred, W., Isarangkul, D., Summpunn, P., Wiyakrutta, S. and Meevootisom, V. (2004) Purification, characterization, gene cloning and sequence analysis of a phytase from Klebsiella pneumoniae subsp. pneumoniae XY-5. ScienceAsia 30, 383–390. Wanner, B.L. (1996) Phosphorus asssimilation and control of the phosphate regulon. In: Neidhardt, F.C., Curtiss, R. III, Ingraham, J.L., Lin, E.C.C., Low, K.B., Magasanik, B., Reznikoff, W.S., Riley, M., Schaechter, M. and Umbarger, H.E. (eds) Escherichia coli and Salmonella: Cellular and Molecular Biology, 2nd edn. ASM Press, Washington, DC, pp. 1357–1381. Whitton, B.A., Graininger, S.L.J., Hawley, G.R.W. and Simon, J.W. (1991) Cell-bound and extracellular phosphatase activities of cyanobacterial isolates. Microbial Ecology 21, 85–98. Yanke, L.J., Bae, H.D., Selinger, L.B. and Cheng, K.-J. (1998) Phytase activity of anaerobic ruminal bacteria. Microbiology 144, 1565–1573. Yoon, S.J., Choi, Y.J., Min, H.K., Cho, K.K., Kim, J.W., Lee, S.C. and Jung, Y.H. (1996) Isolation and identification of phytase-producing bacterium, Enterobacter sp. 4, and enzymatic properties of phytase enzyme. Enzyme and Microbial Technology 18, 449–454. Yoshida, K., Kuromitsu, Z., Ogawa, N. and Oshima, Y. (1989) Mode of expression of the positive regulatory genes PHO2 and PHO4 of the phosphatase regulon in Saccharomyces cerevisiae. Molecular General Genetics 217, 31–39. Zamudio, M., González, A. and Medina, J.A. (2001) Lactobacillus plantarum phytase activity is due to non-specific acid phosphatase. Letters in Applied Microbiology 32, 181–184. Zamudio, M., González, A. and Bastarrachea, F. (2002) Regulation of Raoultella terrigena comb.nov. phytase expression. Canadian Journal of Microbiology 48, 71–81. Zhou, D., Chen, L.-M., Hernandez, L., Shears, S.B. and Galán, J.E. (2001) A Salmonella inositol polyphosphatase acts in conjunction with other bacterial effectors to promote host cell actin cytoskeleton rearrangements and bacterial internalization. Molecular Microbiology 39, 248–259.

7

Phytases: Attributes, Catalytic Mechanisms and Applications Edward J. Mullaney and Abul H.J. Ullah

United States Department of Agriculture–Agricultural Research Service, Southern Regional Research Center, 1100 Robert E. Lee Blvd, New Orleans, LA 70124, USA

Discovered more than a century ago (Posternak, 1903), myo-inositol hexakisphosphate is a ubiquitous constituent in cereals and grains, where it exists predominantly in salt form (phytate). This means that it represents an immense reservoir of phosphorus that can be potentially utilized by plants, microorganisms and animals. Lott et al. (2000) estimated that 51 million t of phytate is sequestered annually in commercially produced crop seeds and fruits. The amount of phosphorus in phytate is therefore equivalent to approximately two-thirds of the phosphorus utilized each year through the application of mineral fertilizers to agricultural land. Phosphorus is an essential component of DNA, adenosine 5′-triphosphate and other compounds necessary for life (Abelson, 1999), so the liberation of inorganic phosphate covalently bound to phytate is essential in numerous organisms. The first phytate-degrading enzyme was reported as early as 1907 (Suzuki et al., 1907) and research continues to the present day. A major impetus for phytase research resulted from the poultry industry switching from fishmeal and other more expensive protein sources to low-cost plant protein such as soybean meal (Rumsey, 1993). Poultry and other animals with simple stomachs lack a digestive phytase, so research to identify a cost-effective phytase that could be added to their diet intensified (Wodzinski

and Ullah, 1996). Today, a number of enzymes that can initiate the cleavage of myo-inositol hexakisphosphate are known to exist in a range of organisms (Konietzny and Greiner, 2002; Simon and Igbasan, 2002; Lei and Porres, 2003; Mullaney and Ullah, 2003; Oh et al., 2004; Haefner et al., 2005; see Hill and Richardson, Chapter 5, and Greiner, Chapter 6, this volume). The detailed characterization of some of these enzymes has revealed that nature did not develop a single catalytic mechanism to cleave phosphate groups from myo-inositol hexakisphosphate in these diverse organisms. In fact, the exact number of catalytic mechanisms that nature has evolved for this purpose is not known. However, it is now clear that different strategies have been adopted to accommodate the physical structure of myo-inositol hexakisphosphate with its six negatively charged phosphate groups as a substrate. The recognition that not all phytases are structurally similar or share a common active site has already yielded an initial classification system based primarily on catalytic mechanism (Oh et al., 2004). However, a taxonomic system needs to be devised that can accommodate new types of phytases with novel catalytic mechanisms. The increasing number of phytate-degrading enzymes now offers scientific investigators the potential to select the catalytic features most amenable to

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their research objectives. Coupled with the potential to engineer certain enzyme characteristics, this offers the potential for significant achievements in this and related fields of research. Today, four distinctly different classes of phosphatase enzymes are known to display phytase activity: 1. 2. 3. 4.

Histidine acid phosphatase (HAP) β-Propeller phytase (BPP) Cysteine phosphatase (CP) Purple acid phosphatase (PAP)

The representatives of each of these classes have different catalytic mechanisms and other unique features that allow them to effectively utilize myoinositol hexakisphosphate as a substrate at various pH values. The fact that each is different also broadens the number of enzymes that may be applied to the search for developing an enhanced phytase for current and future applications of such enzymes.

Histidine Acid Phosphatase HAP is a large class of enzymes with representatives in animals, plants and microorganisms (Wodzinski and Ullah, 1996; Mullaney et al., 2000; Konietzny and Greiner, 2002; Lei and Porres, 2003). A shared characteristic is a common catalytic mechanism. The amino acid RHGXRXP

sequences that compose the active site have been identified and contain an N-terminal motif RHGXRXP and a C-terminal motif HD. When properly folded, these distant sequences converge to form a single catalytic centre, which initiates a two-step reaction that hydrolyses phosphomonoesters (Ullah et al., 1991; van Etten et al., 1991). In this reaction, the histidine residue in the conserved motif of the N-terminal region serves as a nucleophile in the formation of a covalent phospho-histidine intermediate (Ostanin et al., 1992; Lindqvist et al., 1994; Oh et al., 2004). The aspartic acid residue of the C-terminal HD then functions as a proton donor to the oxygen atom of the scissile phosphomonoester bond (Lindqvist et al., 1994; Porvari et al., 1994). The N-terminal active-site motif and the adjacent amino acids found in a wide array of HAPs are presented in Fig. 7.1. It should be noted that the exact sequence is even conserved in the prokaryotic example, Escherichia coli phytase, shown in this figure. It must also be noted that while these enzymes share a common catalytic site, they do not share an equal ability to degrade myo-inositol hexakisphosphate. Both the mouse and fruit fly multiple inositol polyphosphate phosphatase (MIPP) in Fig. 7.1 represent a number of other HAPs that are not effective phytases. Also, the fact that maize phytase displays only 60% homology with the HAP consensus motif (Maugenest et al., 1999), yet is still a phytase, indicates that the ability to utilize myo-inositol

HAP consensus sequence

QVLSRHGARYPTSK

Aspergillus niger NRRL 3135 PhyA

QVLARHGARSPTDS

Aspergillus terreus PhyA

QVLSRHGARYPTES

Aspergillus nidulans PhyA

VIVSRHGVRAPTKA

Escherichia coli phytase (appA gene product)

VALIRHGTRYPTTK

Mus musculus, multiple inositol polyphosphate phosphatase (MIPP)

MWIFRHGDRTPKKS

Drosophila melanogaster MIPP

ELVRRHQLRLGYGS

Maize Phyt I

Fig. 7.1. The conserved N-terminal active site and flanking amino acid sequence in representative histidine acid phosphatases from microorganisms, animals and plants. All enzymes shown, except mouse and Drosophila MIPP (multiple inositol polyphosphate phosphatase), display phytase activity.

Phytases: Attributes, Catalytic Mechanisms and Applications

hexakisphosphate as a substrate depends on more than a single catalytic feature. In recognizing the fact that all HAPs are not phytases, Oh et al. (2004) advanced the term ‘histidine acid phytase’ (HAPhy) to denote HAPs that can accommodate myo-inositol hexakisphosphate as a substrate. Both prokaryotic and eukaryotic HAPhy enzymes are known and share little sequence homology other than the conserved active-site motif. The E. coli phytase is the best-characterized prokaryotic HAPhy (Greiner et al., 1993) and a three-dimensional molecular model of its structure is available (Lim et al., 2000). This enzyme has been advanced for use as an animal feed additive. The successful expression of the E. coli phytase gene in the salivary glands of mice (Golovan et al., 2001a) and swine (Golovan et al., 2001b) was reported recently. The swine, termed the Enviropig™, secretes phytase in its saliva to break down phytate in its feed and produce low-phosphorus manure. Substrate specificity site In eukaryotes, HAPhys have been cloned in maize and a number of fungal isolates. The most widely studied fungal phytases are from Aspergillus niger and A. fumigatus. The crystal structure of both of these enzymes has been derived and deposited in the National Center for Biotechnology Information (NCBI) as 1IHP and 1SK8, respectively. Structural characterization (Kostrewa et al., 1999; Liu et al., 2004; Xiang et al., 2004) and catalytic studies (Wyss et al., 1999a) have identified a new site in the enzyme that facilitates its interaction with different substrates. Kostrewa et al. (1999) identified a region, the substrate specificity site, of the A. niger PhyA molecule that encircles the active site of the enzyme and functions as a ‘gatekeeper’. The same site allows for the interaction of the catalytic site and the highly negatively charged myoinositol hexakisphosphate molecule. In the substrate specificity site of A. niger NRRL 3135 there are two acidic and four basic amino acid residues, E228, D262, K91, K94, K300 and K301 (Kostrewa et al., 1999; Mullaney et al., 2000). This means that at pH 2.5 the four basic amino acids – K91, K94, K300 and K301 – are all positively charged and would attract myo-

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inositol hexakisphosphate. When the pH is raised to 5.0, the local electrostatic field of the substrate specificity site still remains attractive for myoinositol hexakisphosphate. Wyss et al. (1999a) had previously observed that despite the catalytic centres for all the known microbial HAPhys being identical, they could be divided into two classes based on substrate specificity. One class has broad substrate specificity but a low specific activity for myo-inositol hexakisphosphate, while the second class has narrow substrate specificity and a high specific activity for myo-inositol hexakisphosphate. An examination of the amino acids composing the substrate specificity site of the fungal HAPhys in this study revealed a correlation between the amino acid residue 300 and the enzyme’s level of specific activity for myoinositol hexakisphosphate (Mullaney et al., 2002). This study also revealed that residue 301 was strongly conserved as lysine (K), while residue 300 varied considerably. The HAPhys cited in Wyss et al. (1999a) with high specific activity for myoinositol hexakisphosphate have either a basic or acidic amino acid residue at 300, while the phytases with low specific activity have a neutral amino acid at that position. Subsequent sitedirected mutagenesis at residue 300 in A. niger NRRL 3135 PhyA established the importance of the lysine residue at that site and the enzyme’s high specific activity for myo-inositol hexakisphosphate (Mullaney et al., 2002). It should be noted that the replacement of amino acids that are not part of the substrate specificity site has also been reported to enhance the catalytic properties of A. fumigatus phytase (Tomschy et al., 2000). Site-directed mutagenesis studies have linked some of the amino acid residues in the substrate specificity site of A. niger NRRL 3135 to its unique pH, with two optima at 2.5 and 5.0 (Ullah and Gibson, 1987). Explanations for this phenomenon range from dismissal as an artefact (Berka et al., 1998) to possible buffer effects (Lehmann et al., 2000). The selection of an appropriate buffer is of course critical in determining the true specific activity any enzyme displays for myo-inositol hexakisphosphate at various pH values. Recent studies in both A. niger NRRL 3135 (Mullaney et al., 2002) and A. fumigatus ATCC 13073 PhyA (Tomschy et al., 2002) have established that the amino acid residues in the substrate specificity site give rise to this unique two-pH optima profile.

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Differences in the substrate specificity site of a second extracellular A. niger phytase, PhyB, can also explain why it has both a different pH optimum and substrate specificity range than A. niger PhyA. PhyB has only been reported in the isolates of A. niger. Although the active form of PhyA is a monomer, PhyB was reported to be either a dimer (Ullah and Cummins, 1987) or a tetramer (Kostrewa et al., 1999). The individual molecules are electrostatically bound together at the N-terminal region. PhyB displays an optimum pH at 2.5, but unlike PhyA it cannot hydrolyse myo-inositol hexakisphosphate at pH 5.0. That is why researchers named it pH 2.5 optimum acid phosphatase when first characterized (Ullah and Cummins, 1987). Subsequent investigations revealed that it could hydrolyse myo-inositol hexakisphosphate at pH 2.5 (Ullah and Phillippy, 1994). Both PhyB and PhyA share identical active-site characteristics of HAPs, but their substrate specificity sites are remarkably different. Kostrewa et al. (1999) identified the substrate specificity site of PhyB to be composed of only two acidic amino acids, D75 and E272. This means that at pH 5.0 the acidic amino acids would be negatively charged, while at pH 2.5 they would be uncharged. All negatively charged substrates, including myo-inositol hexakisphosphate, would therefore be repelled at the higher pH, but not at the lower. This also explains the substrate specificity difference between PhyA and PhyB. The latter can accept a broader variety of phosphomonoesters, because its substrate specificity site has a more neutral electrostatic field, whereas A. niger’s PhyA has a highly positive electrostatic field at its substrate specificity site and is thus optimized for the binding of negatively charged myo-inositol hexakisphosphate. All the recent findings about the substrate specificity site of fungal HAPs suggest that it has a significant role in determining how effectively the enzyme can hydrolyse myo-inositol hexakisphosphate. By occupying positions adjacent to the catalytic domain, the amino acids in the substrate specificity site function as gatekeepers in determining the ease with which any substrate can pass and interact with the active-site residues. Research is also showing that techniques such as site-directed mutagenesis of a cloned phytase gene can be employed to alter the composition of the enzyme’s substrate specificity site and thus alter both its pH profile and substrate selectivity.

Glycosylation The desire to obtain a commercially viable phytase has focused most research on the isolation of extracellular or secreted enzymes. Consequently, all the fungal phytases that have been characterized thus far are secreted glycoproteins. Glycosylation, the process that adds polysaccharides to proteins, is generally thought to confer stability and assist in the correct folding of the enzyme. The SDS–PAGE profile of purified PhyA provided the first clue that it was heavily glycosylated (Ullah and Gibson, 1987). Sugar analysis of purified phytase indicated the presence of N-glycosidic linkage of high mannose sugar chain to the asparagines (N) (Ullah, 1988). This conclusion was bolstered during chemical sequencing of phytase (Ullah and Dischinger, 1993), which revealed several blank residues linked to the presence of a consensus sequence, or sequon, NXS in the PhyA sequence (Apweiler et al., 1999). The presence of extensive glycosylation on fungal phytase impeded its crystallization for structural studies. It was not until Grueninger-Leitch et al. (1996) developed a recombinant fusion protein glycosidase expressed in E. coli that it became feasible to obtain the high-quality protein crystals that were required to determine a proper three-dimensional X-ray structure of glycoproteins. A. niger PhyA was one of the first enzymes deglycosylated by this technique. All ten N-glycosylation sites of A. niger NRRL 3135 PhyA are indicated in Fig. 7.2. Patterns of glycosylation vary in phytase when expressed in other fungal expression systems and transgenic plants (Ullah et al., 1999, 2002; Wyss et al., 1999b). However, the activity is maintained, and PhyA has been successfully expressed in several fungal expression systems and a number of plant species (Mullaney et al., 2000). Differences in the glycosylation pathway in fungi and animals may explain the low success rate in the expression of any fungal phytase in animals (Bretthauer, 2003). To date, A. niger phy A has been reported to be expressed in only two animals: a fish, the Japanese medaka, Oryzias latipes (Hostetler et al., 2003), and silkworm, Bombyx mori (Wang et al., 2003). However, higher success rates have been obtained with the unglycosylated prokaryotic E. coli phytase, which has been expressed in both mice and swine (Golovan et al., 2001a,b).

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# # MGVSAVLLPLYLLSGVTSGLAVPASRNQSSCDTVDQGYQCFSETSHLWGQYAPFFSLANE

60

# # SVISPEVPAGCRVTFAQVLSRHGARYPTDSKGKKYSALIEEIQQNATTFDGKYAFLKTYN

120

YSLGADDLTPFGEQELVNSGIKFYQRYESLTRNIVPFIRSSGSSRVIASGKKFIEGFQST

180

# # KLKDPRAQPGQSSPKIDVVISEASSSNNTLDPGTCTVFEDSELADTVEANFTATFVPSIR

240

QRLENDLSGVTLTDTEVTYLMDMCSFDTISTSTVDTKLSPFCDLFTHDEWINYDYLQSLK

300

# # KYYGHGAGNPLGPTQGVGYANELIARLTHSPVHDDTSSNHTLDSSPATFPLNSTLYADFS

360

# # HDNGIISILFALGLYNGTKPLSTTTVENITQTDGFSSAWTVPFASRLYVEMMQCQAEQEP

420

LVRVLVNDRVVPLHGCPVDALGRCTRDSFVRGLSFARSGGDWAECFA Fig. 7.2. The amino acid sequence of Aspergillus niger NRRL 3135 PhyA. All ten N-glycosylation sites are indicated by #.

Disulphide bridges Although not directly involved in the catalytic function of HAPhy, disulphide bridges perform an important role in maintaining the proper three-dimensional structure to allow for catalytic activity in phytase (Ullah and Mullaney, 1996; Kostrewa et al., 1997; Wang et al., 2004). The three-dimensional models of A. niger and A. fumigatus PhyA show that all ten cysteine residues present in each enzyme are involved in the formation of five disulphide bridges. An analysis of the deduced amino acid sequence of a number of other fungal phytases that have not been so extensively characterized revealed a pattern of conservation of the cysteines necessary for the formation of these disulphide bridges (Mullaney and Ullah, 2005). A shared eight-cysteine motif (8CM) is widely conserved in all the fungal HAPhys surveyed. The reason for the conservation of this 8CM appears to parallel the recent discovery of another eight-cysteine motif in a number of plant proteins (Jose-Estanyol et al., 2004). In none of these nearly 500 plant polypeptides were any of the cysteines involved in the functional catalytic mechanism of the molecules. Instead, they were conserved to form a network of disulphide bonds that allow for the convergence of sequences nec-

essary for the proper molecular architecture required for folding and specific function of the proteins. In fungal HAPhys, the conservation of the 8CM sequence was 100%, while the overall homology for the sequences examined ranged between 23% and 66%. The survey included amino acid sequences from basidiomycete (Lassen et al., 2001), unicellular and filamentous ascomycete HAPhys (Mullaney and Ullah, 2005). This study also reported that two extra cysteines, which form a fifth disulphide bridge in the Nterminal region of A. niger PhyA, are conserved in all filamentous ascomycete HAPhys. This suggests that the higher stability found in phytases from some Aspergillus spp. may in some manner be correlated with this extra disulphide bridge. In E. coli phytase, all eight cysteines are involved in four disulphide bonds (Lim et al., 2000). However, in this phytase, significantly enhanced activity was achieved when one disulphide bridge was abolished. This was accomplished by site-directed mutagenesis, which replaced a cysteine with another amino acid residue (Rodriguez et al., 2000). It was suggested that the removal of this disulphide bond could modulate the domain flexibility and thereby increase the catalytic efficiency of the enzyme.

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Applications Today, the major application for HAPhys is in the hydrolysis of phytate in cereal grains of animal feed. Numerous feed trials involving poultry, swine and aquaculture have established the efficacy of commercially produced phytase to increase the utilization of phytate while reducing phosphorus concentrations in manure (Kornegay, 1996; Wodzinski and Ullah, 1996; Mullaney et al., 2000). Phytase is now extensively employed worldwide for this application, which is discussed in detail elsewhere in this volume (Lei and Porres, Chapter 9). Food applications include the production of phytate-free soybean milk, low-phytin bread (Dvorakova, 1998; Brinch-Pedersen et al., 2003; Drakakaki et al., 2005) and the production of transgenic grains expressing fungal phytase (Lucca et al., 2001). One novel application was the incorporation of vanadium into the active site of A. niger PhyA to produce a cost-effective semisynthetic peroxidase (van de Velde et al., 2000). A limited market also exists for the use of HAPhy in the production of myo-inositol phosphates. Greiner and Konietzny (1996) demonstrated that a packed-bed bioreactor containing covalently attached E. coli PhyA could cost-effectively produce specific isomers of the lower myo-inositol phosphate esters. HAPhys have been successfully expressed in crop plants demonstrating the potential for ‘biofarming’. The goal in these studies was to prove the feasibility of utilizing transgenic crop plants to produce several enzymes that are currently manufactured at conventional fermentation facilities. To document that this was a cost-effective means to produce bulk enzymes, A. niger phy A was successfully expressed in the fodder crop Medicago sativa (lucerne) (Ullah et al., 2002). Future applications of HAPhys extend to the potential development of plant cultivars that require less phosphorus fertilizer. This research has been prompted by the realization that global phosphate reserves are limited and that future generations may face a ‘phosphate crisis’ (Abelson, 1999). To respond to this challenge, the A. niger phy A gene has been expressed in the roots of a plant, Arabidopsis (Mudge et al., 2003). These transgenic plants can grow on phytate as the sole phosphorus source, which opens up the possibility that crop plants developed from this tech-

nique would require less phosphorus fertilizers and thus slow the depletion of the known supply of phosphorus. To determine the feasibility of this in an agricultural crop, George et al. (2004) engineered a transgenic Trifolium subterraneum L. that exudes an extracellular A. niger PhyA from its roots. This technology is discussed elsewhere in this volume (Richardson et al., Chapter 15).

b-Propeller Phytase Unlike HAPhys, which are members of a wellstudied class of enzymes, β-propeller phytase (BPPhy) represents an entirely new class of enzymes and exhibits no homology to any known phosphatases (Kerovuo et al., 1998; Kim et al., 1998a,b; Ha et al., 2000). The name was adopted for this group of enzymes based on their molecular structure, which consists mainly of β-propeller sheets and resembles a six-bladed propeller (Ha et al., 2000; Shin et al., 2001). Before this, it was termed PhyC (Kerovuo et al., 1998) and TS-Phy (Ha et al., 1999) and has been subsequently labelled PhyD (alkaline phytase) (Oh et al., 2004), PhyL (Tye et al., 2002) and PhyA (Chatterjee et al., 2003). Initially, BPPhys were reported from Bacillus and related bacterial species that require calcium ions for both catalytic activity and thermostability. The calcium facilitates the binding of myo-inositol hexakisphosphate by generating a favourable electrostatic environment in the substrate-binding domain of the biocatalyst. Kinetic studies established that BPPhys could hydrolyse calcium-phytate between pH 7.0 and 8.0 (Oh et al., 2001). The main components involved in the catalytic mechanism of BPPhys to hydrolyse myo-inositol hexakisphosphate include a ‘cleavage site’ and an ‘affinity site’ (Shin et al., 2001). In this model, it is necessary for two adjacent phosphate groups to occupy both the cleavage and affinity sites. The phosphate bound to the affinity site facilitates the cleavage of flanking phosphate by the cleavage site. The enzyme prefers hydrolysis of every second phosphate and has a reduced affinity for any substrate that cannot accommodate this stringent requirement. This explains why BPPhys alternately remove phosphate groups with the end product being myo-inositol trisphosphate. Further degradation then occurs slowly

Phytases: Attributes, Catalytic Mechanisms and Applications

because a neighbouring phosphate group is lacking and the enzyme is increasingly susceptible to product inhibition. Based on its narrow substrate range, a requirement for calcium for catalytic activity, and myo-inositol trisphosphate being the predominant product from myo-inositol hexakisphosphate hydrolysis, Oh et al. (2004) proposed that alkaline phytases from plants share a similar catalytic mechanism with BPPhys. For this reason, several alkaline plant phytases were grouped in the same class with BPPhys. Calcium is known to enhance the activity of several plant alkaline phytases, among others from lily (Lilium longiflorum) pollen (Scott and Loewus, 1986) and a number of legumes (Scott, 1991). Unfortunately, none of the genes thus far have been cloned and no sequence data exist to confirm that they indeed are BPPhys. However, it should be noted that a recent study questions this classification by reporting the presence of the HAP amino acid motif in lily pollen phytase (Mehta and Murthy, 2005). Another similarity has also been noted between BPPhys and pyrophosphatases, which hydrolyse inorganic pyrophosphate. β-Propeller phytases share no homology with any other known classes of phosphatases, which led Hamelryck (2003) to employ a ‘multidimensional index tree’ method to analyse side-chain patterns found in different classes of enzymes. The results suggested that BPPhys and pyrophosphatases share some common structural features, including a cleavage site where the nucleophilic attack by a water molecule transpires, and an affinity site that binds a second phosphate group (Shin et al., 2001). Although BPPhys and pyrophosphatases share a similar catalytic mechanism, the molecular architecture displayed in BPPhys is also found in a number of other proteins. None of these other enzymes are phytases, but several of their β-propeller domains are associated with pathogenesis in various diseases ranging from Alzheimer’s and arthritis to microbial infections (Pons et al., 2003). It is interesting to note that the only phytase associated with pathogenesis to date is a BPPhy. It was reported that Xanthomonas oryzae, a plant pathogen of rice, secretes a sixbladed β-propeller protein, PhyA, which is required for optimum virulence in its host (Chatterjee et al., 2003). Characterization of this protein revealed conservation of active-site

103

residues previously identified in Bacillus phytases. Isolates of X. oryzae having mutant phy A genes display a reduced virulence in their host plants. This is the first study to suggest that virulence in a plant pathogen is due at least in part to the ability of the bacteria to utilize myo-inositol hexakisphosphate in the plant host as a source of phosphate. Bacterial leaf blight caused by X. oryzae is a major rice disease and this raises the possibility that other bacteria and fungal pathogens of plants have evolved a similar or different phytase to access myo-inositol hexakisphosphate in their host. The naming of this BPPhy, also as PhyA, creates the potential for confusion with the HAPhy PhyA and reinforces the need to update the phytase nomenclature. β-Propeller phytases have been advocated for several applications. Their heat tolerance (Kim et al., 1998a) means that they would withstand feed-pelleting, which has made them candidates for use as an animal feed additive, both alone and with HAPhy (Park et al., 1999). Research to advance this has resulted in its successful expression in E. coli to increase its costeffectiveness (Kim et al., 1998b). Bacillus phytase stimulated the growth of maize seedlings under conditions of phosphate limitation but not in the presence of myo-inositol hexakisphosphate (Idriss et al., 2002). Bacillus subtilis phytase has also been expressed in the cytoplasm of transgenic tobacco (Yip et al., 2003). Results indicated that a shift in the equilibrium of the inositol phosphate biosynthesis pathway occurred, which improved plant performance under phosphate starvation. Although these studies offer new strategies for animal feed supplements and possible tools for raising productivity in agriculture, no commercial applications of BPPhys are currently available.

Cysteine Phosphatase Another class of phytase has been reported from an anaerobic ruminal bacterium, Selenomonas ruminantium. It had long been suspected that the reason ruminants could utilize phytate and monogastric animals could not was due in part to the presence of certain microorganisms in the rumen. A survey of anaerobic rumen bacteria revealed phytase activity in one isolate, S. ruminantium (Yanke et al., 1999). Initial characterization

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established that the enzyme was monomeric, ~46 kDa in size, had an optimal pH range of 4.0–5.5, an optimal temperature of 50–55ºC and was inhibited by cations of iron and several other metals. The gene encoding this phytase has now been cloned and its product extensively analysed (Chu et al., 2004). Its crystal structure reveals that it is neither an HAPhy nor a BPPhy. Instead, its structure and proposed catalytic mechanism suggest that it is a member of the (CP) superfamily. It shares the active-site motif HCXXGXXR(T/S) and other substantial similarities with protein tyrosine phosphatase (PTP), a member of the CP group. The active site forms a loop that functions as a substrate-binding pocket unique to PTPs. The depth of this pocket is important because it appears to determine the substrate specificity (Denu and Dixon, 1998). Consistent with this model, the S. ruminantium phytase, the cysteine phytase (CPhy), has a wider and deeper pocket than PTP and is thus able to accommodate the fully phosphorylated inositol group of myo-inositol hexakisphosphate (Chu et al., 2004). The initial binding of myo-inositol hexakisphosphate to the CPhy active-site pocket is facilitated by the negatively charged substrate. The hydrolysis of phosphate groups proceeds sequentially with the end product being myoinositol 2-monophosphate (Chu et al., 2004). The inhibitory effect of iron and other metal cations (copper, zinc and mercury) was attributed to their ability to complex with myo-inositol hexakisphosphate, but the stimulatory effect of lead cations remains unexplained (Yanke et al., 1999). The source of this enzyme, a ruminal bacterium, suggests that it may have evolved many attributes that would lead to its adoption as an animal feed additive. However, at this time no feed trial study or any other commercial applications of CPhy are available.

Purple Acid Phosphatase The PAPs have representatives in plants, mammals, fungi and bacteria (Schenk et al., 2000). Like other metalloenzymes, the active site of PAPs requires one or more metal ions for activity. PAPs share a pattern of five common consensus motifs (DxG/GDx2Y/GNH(E,D)/Vx2H/GHxH) con-

taining seven residues capable of forming metal ligands (Schenk et al., 2000). The first binuclear metal-containing hydrolase identified as a phytase was reported in the cotyledons of a germinating soybean (Glycine max L. Merr.) seedling (Hegeman and Grabau, 2001). The gene encoding GmPhy has been cloned and its product characterized. However, unlike HAPhys, BPPhys and CPhys, no X-ray crystallography study has been performed on PAP phytase (PAPhy) and its three-dimensional structure is unknown. An A. niger PAP (Apase6) was isolated and cloned (Ullah and Cummins, 1988; Mullaney et al., 1995). It does not effectively utilize myoinositol hexakisphosphate as a substrate and there has been no commercial interest in this acid phosphatase. Comparison of the active sites of this A. niger PAP and the soybean PAPhys (Fig. 7.3) indicates that they both contain the conserved active-site motif (Mullaney and Ullah, 1998). When compared to A. niger PhyA, the specific activity for myo-inositol hexakisphosphate of GmPhy is low. The lower catalytic activity of GmPhy may be advantageous during germination because the process requires a steady hydrolysis of myo-inositol hexakisphosphate over this entire period (Mullaney and Ullah, 2003). Chiera et al. (2004) expressed the GmPhy gene in the soybean seed during development, a time when it is not normally expressed. Linking the GmPhy gene to an embryo-specific promoter, β-conglycinin, ectopic expression was achieved and a lower level of phytate occurred in the transformed seeds. A reduction in phytate of up to 25% was achieved. This offers another potential strategy for the development of plant cultivars with lower phytate levels in their seed. Since the reporting of GmPhy, putative PAPhys have been reported in the rice (Oryza sativa L.) genome and cloned from a legume, barrel medic (Medicago. truncatula Gaertn) (Xiao et al., 2005). The latter example revealed high sequence similarity to GmPhy, but unlike the soybean phytase, it does not have all five conserved blocks of the amino acids capable of forming metal ligands that are characteristic of PAP (Xiao et al., 2005). Only limited characterization of M. truncatula PAPhy is currently available, but it has been engineered to be under the control of a root-specific promoter, MtPT1, and expressed in Arabidopsis. The trans-

Phytases: Attributes, Catalytic Mechanisms and Applications

Source

Accession no.

A. niger G. max

Consensus motif

105

* DXG

* * GDXXY

* GNH(E/D)

* VXXH

* * GHXH

JN0656

DMG

GDLSY

GNHE

VLMH

GHIH

AF272346

DLG

GDVTY

GNHE

VTWH

GHVH

Fig. 7.3. Comparison of the amino acids composing the active-site consensus motif of Aspergillus niger NRRL 3135 Apase and Glycine max GmPhy. Asterisks indicate the seven residues in the active site capable of forming ligands with metals. (From Schenk et al., 2000.)

genic phytase was then secreted into the plant’s rhizosphere. This permitted the transgenic Arabidopsis to grow when phytate was the sole source of phosphorus, as reported earlier with the A. niger HAPhy gene expressed in Arabidopsis (Richardson et al., 2001). Xiao et al. (2005) suggested that while both genes could be employed to improve phosphate acquisition in crops, a plant phytase gene could have advantages over a fungal gene with regard to regulatory and biosafety concerns. It was also suggested that it would be worth investigating if the simultaneous expression of the MtPHY1 and a fungal phytase gene in the roots of transgenic plants might benefit phosphorus uptake. Although no PAPhys are currently being marketed, they have been the subject of several research studies.

A Revised Nomenclature for the Phytases Many researchers in the field agree that the expanding interest in phytase over the last decade has created the need to consider a revised nomenclature that more accurately describes the catalytic mechanisms of the numerous enzymes that are currently grouped simply as phytase. Today, the term phytase is primarily based on the in vitro capability of an enzyme to degrade myo-inositol hexakisphosphate (Konietzny and Greiner, 2002; see Greiner, Chapter 6, this volume). It does not identify the catalytic mechanism that is employed by the enzyme to hydrolyse the substrate. Although there are a number of phytases for which the mechanisms of myo-inositol hexakisphosphate hydrolysis are yet to be deter-

mined, the number for which this information is available is steadily increasing (Mullaney and Ullah, 2003). It is thus logical to incorporate the results of this research in a more informative classification system. Differences in the physical features of phytases stem from molecular properties, and Oh et al. (2004) proposed a phytase classification system based on this fact. Physical characteristics of these enzymes, such as pH optima, mineral requirements, substrate requirements and end product of hydrolysis, are all manifestations of the distinct catalytic mechanisms. Different phosphatases have evolved to hydrolyse myo-inositol hexakisphosphate under diversified conditions. Table 7.1 outlines a classification for phytases based on mechanistic enzymology. This classification scheme continues the nomenclature developed by Oh et al. (2004), but incorporates the two classes of phytases, PAPhys and CPhys, that were not included in their system. Although a large number of HAPs, PAPs and CPs cannot degrade myo-inositol hexakisphosphate effectively, members of each class do share a common enzymatic pathway with other members that hydrolyse myo-inositol hexakisphosphate efficiently. Basing this system on enzyme family therefore allows the incorporation of pertinent information developed by research on a significantly larger number of enzymes. This classification system also offers the potential of assigning other uncharacterized phytases based on unique features associated with individual enzyme families. The system is open-ended in that new groups of phytases can easily be added and existing groups subdivided when desirable.

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Table 7.1. Classes of phytases. Organisms in which enzyme occurs

RHGXRXP ... HD

Fungi, plants, bacteria Bacteria (Plant alkaline phytases?) Bacteria

Class

Abbreviation

Enzyme family

Histidine acid phytase β-Propeller phytase

HAPhy BPPhy

Histidine acid phosphatase New family

Cysteine phytase

CPhy

Cysteine phosphatase

HCXXGXXR(T/S)

Purple acid phosphatase

DXG/GDXXY/GNH (E,D)/VXXH/GHXH

Purple acid phytase PAPhy

Six-bladed βpropeller structure

Plants

Unique characteristics

NCBI 3-D model number

Acid phosphatase; EDTA stimulation; final product IP1 Neutral to alkaline phosphatase; calcium required; final product IP3

1IHP 1QFX 1DKP 1H6L

Acid phosphatase; inhibited by Fe2+, Cu2+, Zn2+ and Hg2+; stimulated by Pb2+; final product inositol 2-monophosphate Metalloenzymes

1U24

None currently deposited

E.J. Mullaney and A.H.J. Ullah

Consensus motif or unique structural feature

Phytases: Attributes, Catalytic Mechanisms and Applications

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Note Mention of a trademark or proprietary product does not constitute a guarantee or warranty by the United States Department of Agriculture and does not imply approval to the exclusion of other products that may also be suitable.

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Oh, B.C., Chang, B.S., Park, K.H., Ha, N.C., Kim, H.K., Oh, B.H. and Oh, T.K. (2001) Calcium-dependent catalytic activity of a novel phytase from Bacillus amyloliquefaciens DS11. Biochemistry 40, 9669–9676. Oh, B.C., Choi, W.C., Park, S., Kim, Y.O. and Oh, T.K. (2004) Biochemical properties and substrate specificities of alkaline and histidine acid phytases. Applied Microbiology and Biotechnology 63, 362–372. Ostanin, K., Harms, E.H., Stevis, P.E., Kuciel, R., Zhou, M.M. and van Etten, R.L. (1992) Overexpression, sitedirected mutagenesis, and mechanism of Escherichia coli acid phosphatase. Journal of Biological Chemistry 267, 22830–22836. Park, S.C., Choi, Y.W. and Oh, T.K. (1999) Comparative enzymatic hydrolysis of phytate in various animal feedstuff with two different phytases. Journal of Veterinary Medical Science 61, 1257–1259. Pons, T., Gomez, R., Chinea, G. and Valencia, A. (2003) Beta-propellers: associated functions and their role in human diseases. Current Medicinal Chemistry 10, 505–524. Porvari, K.S., Herrala, A.M., Kurkela, R.M., Taavitsainen, P.A., Lindqvist, Y., Schneider, G. and Vihko, P.T. (1994) Site-directed mutagenesis of prostatic acid phosphatase. Catalytically important aspartic acid 258, substrate specificity, and oligomerization. Journal of Biological Chemistry 269, 22642–22646. Posternak, M.S. (1903) Sur un nouveau principe phosphor-organique d’origine vegetale, la phytine. Comptes Rendus des Séances de la Societe de Biologie et de Ses Filiales 55, 1190–1192. Richardson, A.E., Hadobas, P.A. and Hayes, J.E. (2001) Extracellular secretion of Aspergillus phytase from Arabidopsis roots enables plants to obtain phosphorus from phytate. The Plant Journal 25, 641–649. Rodriguez, E., Wood, Z.A., Karplus, A. and Lei, X.G. (2000) Site-directed mutagenesis improves catalytic efficiency and thermostability of Escherichia coli pH 2.5 acid phosphatase/phytase expressed in Pichia pastoris. Archives of Biochemistry and Biophysics 382, 105–112. Rumsey, G.L. (1993) Fishmeal and alternate sources of protein in fish feeds: update 1993. Fisheries 18, 14–19. Schenk, G., Guddat, L.W., Ge, Y., Carrington, L.E., Hume, D.A., Hamilton, J. and de Jersey, J. (2000) Identification of mammalian-like purple acid phosphatases in a wide range of plants. Gene 250, 117–125. Scott, J. J. (1991) Alkaline phytase activity in nonionic detergent extracts of legume seeds. Plant Physiology 95, 1298–1302. Scott, J. J. and Loewus, F.A. (1986) A calcium-activated phytase from pollen of Lilium longiflorum. Plant Physiology 82, 333–335. Shin, S., Ha, N.C., Oh, B.C., Oh, T.K. and Oh, B.H. (2001) Enzyme mechanism and catalytic property of β propeller phytase. Structure 9, 851–858. Simon, O. and Igbasan, F. (2002) In vitro properties of phytases from various microbial origins. International Journal of Food Science and Technology 37, 813–822. Suzuki, U., Yoshimura, K. and Takaishi, M. (1907) Ueber ein Enzym ‘Phytase’ das ‘Anhydro-oxy-methylen diphosphorsaure’ spaltet. Tokyo Imperial University College of Agriculture Bulletin 7, 503–512. Tomschy, A., Tessier, M., Wyss, M., Brugger, R., Broger, C., Schnoebelen, L., van Loon, A.P.G.M. and Pasamontes, L. (2000) Optimization of the catalytic properties of Aspergillus fumigatus phytase based on the three-dimensional structure. Protein Science 9, 1304–1311. Tomschy, A., Brugger, R., Lehmann, M., Svendsen, A., Vogel, K., Kostrewa, D., Lassen, S., Burger, D., Kronenberger, A., van Loon, A.P.G.M., Pasamontes, L. and Wyss, M. (2002) Engineering of phytase for improved activity at low pH. Applied and Environmental Microbiology 68, 1907–1913. Tye, A. J., Siu, F.K., Leung, T.Y. and Lim, B.L. (2002) Molecular cloning and the biochemical characterization of two novel phytases from B. subtilis 168 and B. licheniformis. Applied Microbiology and Biotechnology 59, 190–197. Ullah, A.H. J. (1988) Aspergillus ficuum phytase: partial primary structure, substrate selectivity, and kinetic characterization. Preparative Biochemistry 18, 459–471. Ullah, A.H. J. and Cummins, B. J. (1987) Purification, N-terminal amino acid sequence and characterizatiom of pH 2.5 optimum acid phosphatase (E.C.3.1.3.2) from Aspergillus ficuum. Preparative Biochemistry 17, 397–422.

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Ullah, A.H. J. and Cummins, B. J. (1988) Aspergillus ficuum extracellular pH 6.0 optimum acid phosphatase: purification, N-terminal amino acid sequence, and biochemical characterization. Preparative Biochemistry 18, 37–65. Ullah, A.H. J. and Dischinger, H.C. Jr (1993) Aspergillus ficuum phytase: complete primary structure elucidation by chemical sequencing. Biochemical and Biophysical Research Communications 92, 747–753. Ullah, A.H. J. and Gibson, D.M. (1987) Extracellular phytase (E.C.3.1.3.8) from Aspergillus ficuum NRRL 3135: purification and characterization. Preparative Biochemistry 17, 63–91. Ullah, A.H. J. and Mullaney, E. J. (1996) Disulfide bonds are necessary for structure and activity in Aspergillus ficuum phytase. Biochemical and Biophysical Research Communications 227, 311–317. Ullah, A.H. J. and Phillippy, B.Q. (1994) Substrate selectivity in Aspergillus ficuum phytase and acid phosphatases using myo-inositol phosphates. Journal of Agricultural and Food Chemistry 42, 423–425. Ullah, A.H. J., Cummins, B. J. and Dischinger, H.C. Jr (1991) Cyclohexanedione modification of arginine at the active site of Aspergillus ficuum phytase. Biochemical and Biophysical Research Communications 178, 45–53. Ullah, A.H. J., Sethumadhavan, K., Mullaney, E. J., Ziegelhoffer, T. and Austin-Phillips, S. (1999) Characterization of recombinant fungal phytase (phy A) expressed in tobacco leaves. Biochemical and Biophysical Research Communications 264, 201–206. Ullah, A.H.J., Sethumadhavan, K., Mullaney, E. J., Ziegelhoffer, T. and Austin-Phillips, S. (2002) Cloned and expressed fungal phyA gene in alfalfa produces a stable phytase. Biochemical and Biophysical Research Communications 290, 1343–1348. van de Velde, F., Konemann, L., van Rantwijk, F. and Sheldon, R.A. (2000) The rational design of semisynthetic peroxidases. Biotechnology and Bioengineering 67, 87–96. van Etten, R.L., Davidson, R., Stevis, P.E., MacArthur, H. and Moore, D.L. (1991) Covalent structure, disulfide bonding, and identification of reactive surface and active site residues of human prostatic acid phosphatase. Journal of Biological Chemistry 266, 2313–2319. Wang, W.B., Yao, B., Xioa, Q.L., Ji, P., Wang, S.P., He, J.L. and Wu, X.F. (2003) Expression of phytase gene in Bombyx mori. Sheng Wu Gong Cheng Xue Bao 19, 112–115. Wang, X.Y., Meang, F.G. and Zhou, H.M. (2004) The role of disulfide bonds in the conformational stability and catalytic activity of phytase. Biochemistry and Cell Biology 82, 329–334. Wodzinski, R.J. and Ullah, A.H.J. (1996) Phytase. Advances in Applied Microbiology 42, 263–302. Wyss, M., Brugger, R., Kronenberger, A., Remy, R., Fimbel, R., Oesterhelt, G., Lehmann, M. and van Loon, A.P.G.M. (1999a) Biochemical characterization of fungal phytases (myo-inositol hexakisphosphate phosphohydrolases): catalytic properties. Applied and Environmental Microbiology 65, 367–373. Wyss, M., Pasamontes, L., Friedlein, A., Remy, R., Tessier, M., Kronenberger, A., Middendorf, A., Lehmann, M., Schnoebelen, L., Rothlisberger, U., Kusznir, E., Wahl, G., Muller, F., Lahm, H.W., Vogel, K. and van Loon, A.P.G.M. (1999b) Biophysical characterization of fungal phytases (myo-inositol hexakisphosphate phosphohydrolases): molecular size, glycosylation pattern, and engineering of proteolytic resistance. Applied and Environmental Microbiology 65, 359–366. Xiang, T., Liu, Q., Deacon, A.M., Koshy, M., Kriksunov, I.A., Lei, X.G., Hao, Q. and Thiel, D.J. (2004) Crystal structure of a heat-resilient phytase from Aspergillus fumigatus, carrying a phosphorylated histidine. Journal of Molecular Biology 339, 437–445. Xiao, K., Harrison, M.J. and Wang, Z. (2005) Transgenic expression of a novel M. truncatula phytase gene results in improved acquisition of organic phosphorus by Arabidopsis. Planta 222, 27–36. Yanke, L.J., Selinger, L.B. and Cheng, K.J. (1999) Phytase activity in Selenomonas ruminantium: a preliminary characterization. Letters in Applied Microbiology 29, 20–25. Yip, W., Wang, L., Cheng, C., Wu, W., Lung, S. and Lim, B.L. (2003) The introduction of a phytase gene from Bacillus subtilis improved the growth performance of transgenic tobacco. Biochemical and Biophysical Research Communications 310, 1148–1154.

8

Seed Phosphorus and the Development of Low-Phytate Crops Victor Raboy United States Department of Agriculture–Agricultural Research Service, Small Grains and Potato Germplasm Research Unit, 1691 S. 2700 W., Aberdeen, ID 83210, USA

Phytate (salts of myo-inositol hexakisphosphate) represents between 60% and 80% of mature seed total phosphorus (Raboy, 1997). Normal or ‘wildtype’ seeds produced by the major cereal crops such as maize (Zea mays L.), wheat (Triticum aestivum L.), barley (Hordeum vulgare L.) or rice (Oryza sativa L.) typically contain between 3.0 and 4.0 mg P/g dry wt (Fig. 8.1, left; Raboy, 1997; Lott et al., 2000). Seeds produced by the major legume soybean (Glycine max L [Merr.]) typically have a higher total phosphorus concentration between 6.0 and 8.0 mg P/g dry wt. In both cases, between 65% and 75% (±10%) of the total phosphorus is found as phytate (illustrated for the cereal crops in Fig. 8.1, left). Inorganic phosphate normally represents about 5% (±3%) of the remaining phosphorus, while lower-order myoinositol phosphates usually represent <10%. Phytic acid, the free-acid form of myo-inositol hexakisphosphate, is ubiquitous in eukaryotes. Its metabolism is important to a number of processes and functions in the eukaryotic cell, ranging from phosphorus and mineral storage in seeds to signal transduction, vesicular trafficking, stress response, RNA transport, DNA metabolism and the regulation of development (reviewed in Raboy, 2003; Shears, 2004). During seed development phytate mostly accumulates as mixed ‘phytin’ salts of several mineral cations. These are primarily mixed potassium and magnesium salts, probably reflecting the relative abundance of these elements in the seed. However, calcium, manganese, zinc and

iron phytates are also found (Lott et al., 1995). Phytate is often deposited as discrete globular inclusions called globoids, located within protein storage vacuoles (PSVs). In cereal grains, starchy endosperm PSVs primarily contain storage protein deposits, whereas phytins are localized in the aleurone and germ (embryo and scutellum) PSVs. In maize, >80% of seed phytate is localized in the germ, with the remainder in the aleurone. In small grains such as wheat, barley and rice, the opposite occurs; ≥80% of seed phytate is localized in the aleurone, with the remainder in the germ (O’Dell et al., 1972). In the soybean seed phytin deposits are dispersed throughout the germ and cotyledonary tissues. The primary applied interest in seed phytate concerns its role in the nutritional quality of feeds and foods prepared from seeds. In the case of animal feeds, the primary interest is in phytate as the seed’s major reserve of phosphorus. Nonruminant (also known as monogastric) animals such as poultry, swine and fish do not efficiently digest and utilize phytate (Brinch-Pedersen et al., 2002), which has two negative outcomes. First, non-ruminant feed must be supplemented with mineral phosphate to provide the animal’s nutritional requirement for phosphorus. Second, the excretion of phytate can be an environmental issue, as phosphorus in animal manure can contribute to water pollution (Sharpley et al., 1994; see Leytem and Maguire, Chapter 10, this volume). Reducing the environmental impact of

©CAB International 2007. Inositol Phosphates: Linking Agriculture and the Environment (eds B.L. Turner, A.E. Richardson and E.J. Mullaney)

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Seed phosphate fraction (mg P/g)

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4

2

0

Wild-type

Ipa1-1

Ipa2-1

Maize genotype Phytate

Inorganic P

Lower inositol P

Other P

Fig. 8.1. Typical seed phosphorus fractions in three types of maize genotypes: homozygous wildtype or ‘normal’; low phytic acid 1-1; low phytic acid 2-1. ‘Lower inositol P’ refers to inositol phosphates with five or fewer phosphate esters per molecule compared with six in phytate. ‘Other P’ refers to all forms of phosphorus in seeds other than in phytate, lower inositol phosphates or inorganic phosphates, such as DNA, RNA, protein and phospholipids.

livestock production remains a major problem for producers and society (Environmental Protection Agency, 2002). One approach to problems associated with feed phytate is to supplement feeds with the enzyme phytase (Brinch-Pedersen et al., 2002; see Lei and Porres, Chapter 9, this volume). As ruminants such as beef and dairy cattle typically digest most feed phytate, it is generally not considered a major issue in this context. In contrast to the relatively straightforward issues concerning feed phytate and phosphorus management in non-ruminant production, their role in human nutrition and health are far more complicated. In the context of human nutrition, consumption of seed-derived phytate can contribute to mineral deficiencies, including iron, zinc, magnesium and calcium deficiencies. This is of particular concern to populations that depend on grains and legumes as staple foods, and especially for women of childbearing age, infants and children within such populations (Brown and Solomons, 1991; Hurrell, 2003). However, dietary phytate may have a positive role as an anti-oxidant and anti-cancer agent (Graf et al.,

1987; Shamsuddin et al., 1988; Vucenik and Shamsuddin, 2003; Singh and Agarwal, 2005; Somasundar et al., 2005) or as an inhibitor of renal stone formation (Grases et al., 2000). These issues remain unresolved and numerous ongoing studies continue to address them. Two observations need to be made concerning the issue of dietary phytate in human nutrition and health. First, the negative impacts of dietary phytate are considered most important in relatively younger people in the developing world who rely on cereal grains and legumes as staple foods. In contrast, the potentially positive roles appear to be most important to the health of ageing individuals in developed countries. Thus, the issue of dietary phytate in human nutrition and health must be considered on a case-by-case basis. Second, in the debate concerning the importance of dietary phytate there is a general lack of cross-communication between the fields of human and animal nutrition. In particular, it often appears that those interested in the issue of dietary phytate in human nutrition and health ignore the issue of dietary phytate in livestock production and the results of the numerous studies on the impact of phytate in animal feed. This is unfortunate, because cereal or legume crop improvement or production is only rarely an issue of producing food for people. Crop species may often be used both in animal feed and human food. The investigation of low phytic acid (lpa) genotypes of major crops began in the early 1990s with the isolation of two maize mutants (Raboy et al., 2000; Fig. 8.1, centre and right). These can be used to develop ‘low-phytate’ crops, which represent a second approach to addressing the dietary and environmental problems associated with seed phytate. In the seeds of these genotypes (Table 8.1), phytate is reduced by between 30% and 90%, but total phosphorus is typically not altered to a great extent (a first exception to this general rule, barley lpa1-1, is discussed below). Instead, reductions in seed phytate are largely matched by increases in inorganic phosphate (Fig. 8.1, lpa1-1), or in some cases by increases in lower-order myo-inositol phosphates, such as myo-inositol tetrakisphosphate or myo-inositol pentakisphosphate. These compounds contain four or five phosphate esters per molecule, as compared with the six phosphates of myo-inositol hexakisphosphate (Fig. 8.1, centre and right).

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Table 8.1. Seed phosphorus fractions in maize, barley, rice, wheat and soybean low phytic acid genotypes.a Species

Genotype

Maize Maize Maize Barley Barley Barley Barley Barley Rice Rice Wheat Wheat Soybean Soybean

Wild-type lpa1-1 lpa2-1 Wild-type lpa1-1 (M 422) lpa2-1 (M 1070) lpa 3-1 (M 635) M 955 Wild-type lpa1-1 Wild-type JS-12-LPA Wild-type M 153

Total P (g P/kg)

Total inositol P

4.5 4.7 4.6 4.8 3.7 5.4 5.0 5.0 3.1 3.5 5.3 5.1 7.95 7.96

3.4 1.1 2.6 2.9 1.2 1.9 0.7 NDc 2.23 1.37 4.0 2.5 5.68 1.98

Inorganic P 0.3 3.1 1.3 0.4 1.2 1.7 2.5 3.3 0.14 1.13 0.5 2.6 0.30 3.02

Cellular Pb 0.8 0.5 0.7 1.5 1.3 1.7 1.8 1.7 0.79 1.05 0.8 NDc 1.97 2.96

a

All analyses listed in this table were conducted in the USDA–ARS Cereal Chemistry (Raboy) laboratory. (From Larson et al., 2000; Raboy et al., 2000; Wilcox et al., 2000; Dorsch et al., 2003; Oltmans et al., 2005 using methods as described.) Total inositol phosphate is primarily myo-inositol hexakisphosphate, but the analytical method used will also detect lower-order inositol phosphates if they are present. Cellular phosphorus includes all forms of phosphorus other than inositol phosphate and inorganic phosphate, including phosphorus in starch, phospholipids, RNA and DNA. b Cellular phosphorus = total phosphorus – (inositol phosphate + inorganic phosphate). c ND = not detected. Inositol phosphate concentrations in M 955 were below that which is reliably assayed with the methods used. In the case of the wheat JS-12-LPA mutant, in the analyses given in this table, the sum of inositol phosphate and inorganic phosphate equalled total phosphorus, indicating no other forms of cellular phosphorus. This is clearly an artefact of this particular assay.

The biochemistry and molecular genetics of phytate metabolism has also continued to progress greatly. Many of the enzymes involved in the phosphorylation of myo-inositol to the hexakisphosphate have been described and genes encoding such enzymes have been identified. In some cases, lpa genotypes have been used in ‘forward genetics’ approaches to identify genes and functions important to seed phytate synthesis (Shi et al., 2003, 2005a). Recent studies using Arabidopsis, the model organism for much plant science research, are also contributing advances to this field (Stevenson-Paulik et al., 2002, 2005). In contrast, relatively little progress has been made in the molecular biology of seed total phosphorus, defined as the sum of all forms of phosphorus in the seed. Manipulating the amount of phytate in seeds, such as in lpa genotypes, has not greatly influenced seed total phosphorus. In lpa genotypes the chemistry of the phosphorus is altered, but not the total concentration. Reduced total phosphorus may have value when seed crops are used in ruminant (dairy and beef) feed ( Volk et al., 2000; Erickson

et al., 2002; Rotz et al., 2002; Toor et al., 2005; see Dao, Chapter 11, this volume), as well as in other applications, including non-ruminant feed. While a great deal of progress has been made in understanding the molecular biology of the uptake and distribution of phosphorus in the parent plant, less is known about the uptake and distribution of phosphorus in the seed. This chapter reviews the biochemistry and genetics of seed phytate and lpa genotypes and the breeding and nutritional evaluation of ‘lowphytate’ crops. It concludes with a look at future directions for research on seed phosphate.

Metabolic Pathways, Genes and Mutants Substantial progress has been made in the molecular genetics and biochemistry of phytic acid biosynthesis, which represents one component of the myo-inositol phosphate pathways. In developing seeds the myo-inositol phosphate metabolic

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of the myo-inositol ring (Loewus and Murthy, 2000). Its activity is coupled with myo-inositol 3-monophosphatase (IMP) in a simple MIPS/IMP ‘myo-inositol synthesis pathway’. Hitz et al. (2002) demonstrated that the LR33 mutation in a soybean MIPS gene resulted in a block in seed phytate accumulation, indicating that a substantial fraction of the myo-inositol necessary for phytate synthesis in

pathways can be viewed as consisting of two parts: (i) synthesis and/or supply of myo-inositol and phosphate (Fig. 8.2, top); and (ii) myo-inositol phosphate/phosphatidylinositol (PtdIns) phosphate metabolism leading to myo-inositol trisphosphates and ultimately phytic acid (Fig. 8.2, middle and bottom). The enzyme D-myo-inositol 3-monophosphate synthase (MIPS) is the sole synthetic source

Glucose 6-phosphate

Soybean LR33 mutation (Hitz et al., 2002)

Ins(3)P1 synthase OH

H HO HO 5

4

H

2

OH

H OH

1

H

HO HO 5

Inositol monophosphatase

H

PtdIns

PtdIns(4)P1 PtdIns(4)P1 5-kinase PtdIns(4,5)P2

Phospholipase C OH

H

P

H

H

H OH

1

P HO

H

4

3 6 1 OH H H H

P 5

Ins(1,3,4,5)P4

Ins(1,3,4)P3

Ins(3,4,6)P3

P

P

5

H

H

2

Inositol polyphosphate kinases H

OH

P

4

H

H

P

3 6 1 OH H H H

H

2 P

Arabidopsis IPK2 mutants (Stevenson-Paulik et al., 2005)

H

2

3 6

5

4

P

H P

OH

Ins(1,4,5)P3

Maize lpa2 (Shi et al., 2003)

Inositol polyphosphate kinases

Phosphatidyl inositol phosphate early intermediate pathway

OH

H

P

H 2

3 6

H

PtdIns 4-kinase

Phosphatidylinositol synthase (PtdIns synthase)

Maize lpa3 (Shi et al., 2005b)

4 P

OH

Inositol kinase

H

5

H

2

myo-Inositol (Ins)

Inositol phosphate early intermediate pathway

HO

4

3 6 H 1 OH H H

Ins(3)P1

P

OH

H

3 6

OH

H

P

H

P 1 H

Ins(1,3,4,5,6)P5

Arabidopsis IPK1 mutants (Stevenson-Paulik et al., 2005)

Inositol polyphosphate 2-kinase P

H P

H P

Breakdown via phytases

4

P 5 H

P

H

H

2

3 6

P 1 H

More highly phosphorylated Ins hepta- and octa-phosphates InsP7 or InsP8

Ins(1,2,3,4,5,6)P6 (phytic acid)

Fig. 8.2. Biochemical pathways leading from glucose 6-phosphate to myo-inositol (Ins) and ultimately to myo-inositol hexakisphosphate. Steps in the pathways for which there are known mutations or blocks are indicated by crossing lines accompanied by the published reference describing the mutation(s). The six carbons of the myo-inositol ring are numbered according to the D-numbering convention. P represents a phosphate group, PH2O4.

Seed Phosphorus and Low-Phytate Crops

seeds is synthesized de novo. Yoshida et al. (1999) had previously demonstrated that MIPS is expressed proximal to the site of phytate accumulation during the development of rice seeds. Pathways to phytic acid might then either proceed through sequential phosphorylation of myo-inositol and ‘soluble myo-inositol phosphates’ (Fig. 8.2, middle-left) or by one of two alternative pathways that utilize PtdIns early intermediates (Fig. 8.2, middle-right). In the pathway proposed by Biswas et al. (1978a), myo-inositol 3-monophosphate is directly converted to myo-inositol 1,3,4,5,6-pentakisphosphate through sequential phosphorylation by a phosphoinositol kinase, of which two electrophoretic forms were identified. The conversion of myo-inositol 1,3,4,5,6-pentakisphosphate to phytic acid is then catalysed by a myo-inositol hexakisphosphate–adenosine diphosphate phosphotransferase (Biswas et al., 1978b), now commonly referred to as myo-inositol 1,3,4,5,6-pentakisphosphate 2-kinase (Phillippy et al., 1994). An alternative pathway to phytic acid begins with myo-inositol as the initial substrate and myo-inositol kinase activity (E.C. 2.7.1.6.4; English et al., 1966; Loewus et al., 1982) and proceeds through site-specific sequential phosphorylation steps of defined soluble myoinositol phosphates (Fig. 8.2, middle-left). This pathway was described in studies of the cellular slime mould Dictyostelium discoideum (Stephens and Irvine, 1990) and the monocot Spirodela polyrhiza (Brearley and Hanke, 1996b). These ‘inositol phosphate early intermediate’ (Fig. 8.2, left side) or ‘lipid-independent’ (Stevenson-Paulik et al., 2002) pathways are similar, and share the common intermediate myo-inositol 3,4,6-trisphosphate (Fig. 8.2). Recently, Shi et al. (2005) determined that the maize lpa3 gene encodes myo-inositol kinase, providing the first genetic evidence for the existence of myo-inositol kinase and demonstrating its importance to phytate synthesis and accumulation in seeds. The pathways to phytic acid that involve PtdIns phosphate lipid intermediates and myoinositol 1,4,5-trisphosphate are illustrated in Fig. 8.2 (middle-right). In a pathway to phytic acid expressed in the nucleus of yeast (York et al., 1999), PtdIns 4,5-bisphosphate is hydrolysed to yield myo-inositol 1,4,5-trisphosphate, which is then phosphorylated directly to myo-inositol 1,3,4,5,6-pentakisphosphate by myo-inositol 1,4,5trisphosphate 3/6-kinase that is encoded by the

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IPK2 gene. A variety of names have been used for this and other types of inositol phosphate kinases, but for simplicity it is referred to here only as myo-inositol 1,4,5-trisphosphate 3/6kinase. Similarly, the second type of multifunctional myo-inositol polyphosphate kinase discussed immediately below is referred to only as myo-inositol 1,3,4-trisphosphate 5/6-kinase. StevensonPaulik et al. (2002) isolated two Arabidopsis thaliana genes related to the yeast myo-inositol 1,4,5trisphosphate 3/6-kinase, termed AtIpk2α and AtIpk2β. Stevenson-Paulik et al. (2005) recently demonstrated that mutations in AtIpk2β impact on the ability of seeds to synthesize phytic acid, indicating that the pathway does proceed at least in part via the intermediate myo-inositol 1,4,5trisphosphate. Alternatively, studies of human cells described a pathway whereby myo-inositol 1,4,5-trisphosphate is first converted to myo-inositol 1,3,4,5-tetrakisphosphate and then to myo-inositol 1,3,4-trisphosphate (Fig. 8.2, centre), which is subsequently phosphorylated to myo-inositol 1,3,4,5,6-pentakisphosphate through what was first defined as myo-inositol 1,3,4trisphosphate 5/6-kinase (Wilson and Majerus, 1996, 1997). The maize lpa2 gene encodes myoinositol 1,3,4-trisphosphate 5/6-kinase (Shi et al., 2003). In seeds produced by plants homozygous for maize lpa2 null mutations, phytic acid is reduced by 35%, clearly indicating that this second type of myo-inositol polyphosphate kinase is important to seed phytic acid synthesis. myo-Inositol 1,3,4trisphosphate and myo-inositol 1,4,5-trisphosphate kinases, such as those encoded by maize lpa2 and Arabidopsis AtIpk2β, respectively, can phosphorylate multiple myo-inositol phosphates (reviewed in Shears, 2004). One or perhaps both types of myoinositol polyphosphate kinases working together might be able to convert myo-inositol 3,4,6-trisphosphate to myo-inositol 1,3,4,5,6-pentakisphosphate. Thus, a pathway to phytate that proceeds entirely via soluble myo-inositol phosphates might in fact utilize enzymes considered part of the PtdIns-intermediate pathway to phytate (Stevenson-Paulik et al., 2002; Raboy, 2003). In summary, it is of interest that the three kinase mutations that perturb phytic acid accumulation in seed (maize lpa3, maize lpa2 and Arabidopsis Ipk2) represent genes encoding functions that represent markers for the three alternative pathways to phytate. Maize lpa3 mutants block myo-inositol kinase activity believed to catal-

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yse the first step in the soluble myo-inositol phosphate/lipid-independent pathway; Arabidopsis IPK2 mutants block myo-inositol 1,4,5-trisphosphate 3/6-kinase activity, critical to the type of lipid-dependent pathway that proceeds directly from myo-inositol 1,4,5-trisphosphate to myo-inositol 1,3,4,5,6-pentakisphosphate; maize lpa2 mutants block myo-inositol 1,3,4-trisphosphate 5/6-kinase activity, believed to be critical to an alternative lipid-dependent pathway first described in the studies of human cells. This indicates that either the pathway to phytic acid in seeds is complex and non-linear or that our understanding and definition of these genes and enzymes requires revision. Regardless of precursor pathways, there is a consensus that myo-inositol 1,3,4,5,6-pentakisphosphate is the penultimate myo-inositol phosphate in the pathway to phytic acid, and that its conversion is catalysed by myoinositol polyphosphate 2-kinase (Biswas et al., 1978a,b; York et al., 1999). Stevenson-Paulik et al. (2005) demonstrated that a mutation in an Arabidopsis gene encoding a 2-kinase substantially blocked seed phytic acid accumulation. The existence of myo-inositol phosphates more highly phosphorylated than phytic acid seems to occur relatively widely in eukaryotic cells (Stephens et al., 1993; Laussmann et al., 2000). These compounds contain pyrophosphate moieties and include 5-diphosphoinositol

1,2,3,4,6-pentakisphosphate (with seven phosphate groups) and bis-diphosphoinositol 1,2,3,4tetrakisphosphate (with eight phosphate groups). To date there has been very little progress in the study of these pyrophosphate-containing compounds in plant systems (Brearley and Hanke, 1996a; Flores and Smart, 2000; Dorsch et al., 2003). Their potential role in seed phytate metabolism, localization or deposition is unknown.

Phytate Deposition in Globoids While this brief review indicates that much progress has been made in the molecular biology of the structural synthetic pathway leading to phytic acid synthesis, relatively little progress has been made regarding how phytate salts are formed and deposited as globoids. Little is known about how the proteins, substrates and products important to these pathways and processes are localized and compartmentalized within the cell. Figure 8.3 provides a model for a phytateaccumulating PSV or membrane-bound globoid, illustrating several of the localization and transport functions of possible importance. An ongoing debate concerns whether or not individual globoids are membrane-bound. Jiang et al.

Fig. 8.3. Components of a membrane-bound organelle, representing either a protein storage vacuole (PSV) or a membrane-bound globoid found within a compound PSV. V-PPase = vacuolar inorganic pyrophosphatase; V-ATPase = vacuolar adenosine triphosphatase; TIP = tonoplast intrinsic protein; PtdIns = phosphatidylinositol. Question marks indicate speculative aspects of the diagram.

Seed Phosphorus and Low-Phytate Crops

(2001) provided evidence that globoids are membrane-bound, and therefore that the PSVs that contain them represent a ‘compound organelle’. For the purpose of this discussion, Fig. 8.3 illustrates an organelle defined by a single membrane. The PtdIns phosphates that might serve as intermediates in the synthesis of phytic acid are themselves localized to the membrane that binds the phytate-accumulating PSV. Specific molecular details of how mixed phytate salts are deposited within seed PSVs are unknown. Water is probably transported into the PSV by aquaporins, one of the several isoforms of tonoplast intrinsic proteins. One or more such proteins might be specific to the globoid containing PSV ( Jauh et al., 1999; Takahashi et al., 2004). Second, the PSV (or globoid) membrane probably contains both vacuolar pyrophosphatases (VPPase) and adenosine triphosphatase, which break down inorganic pyrophosphate or adenosine triphosphate, respectively, and pump protons into the internal vacuolar space (Maeshima, 2000; Jiang et al., 2001). This establishes a proton gradient that drives transport, through channels and antiporters, of various solutes from the cytoplasm into the PSV. These solutes must include potassium and magnesium counter-ions, and possibly other minerals. Adenosine triphosphate–binding cassette transporters (ABCs), which use the energy provided by adenosine triphosphate breakdown to transport a variety of solutes ( Jasinski et al., 2003), play some role since maize lpa1 encodes an ABC transporter (Shi et al., 2005a). As transport functions are probably important to seed phytate deposition, mutations in genes encoding such functions probably would impact net seed phytate accumulation. The first example of a single-gene mutation is a transport function that impacts seed phytate is Maize lpa1 (Shi et al., 2005a). Quantitative variation in the levels of phosphate and phytate in vegetative and seed tissues of Arabidopsis was used to identify a quantitative trait locus (QTL) that accounts for a significant amount of the variation observed (Bentsink et al., 2003). Contained within the 99kb chromosomal segment represented by this QTL were 13 open reading frames, one of which encoded a putative vacuolar adenosine triphosphatase (V-ATPase). Bentsink et al. (2003) hypothesized that the variation in phosphate and phytate levels observed among the Arabidopsis lines in their study was in large part due to varia-

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tion in phosphate transport caused by heritable differences in this enzyme.

Low-Phytate Crops The first two lpa mutations were isolated in maize (maize lpa1-1 and maize lpa2-1; Fig. 8.1; Raboy et al., 2000) by screening seed sampled from chemically mutagenized populations directly for reduced seed phytic acid, using a paper electrophoresis method. Analyses of seed produced by plants homozygous for either maize lpa1-1 or lpa2-1 indicated that the reductions in phytic acid were largely matched by increases in inorganic phosphate, so that total phosphorus in the seed remained unchanged and similar to the wild type (Fig. 8.1; Table 8.1). The increase in seed inorganic phosphate in these first two lpa mutants was several-fold, from less than 0.5 mg P/g in nonmutant seed to between 1.0 and 3.0 mg P/g in the mutants. As accurate tests for seed inorganic phosphate are much quicker and straightforward than methods to assay phytic acid (such as those requiring precipitation, purification or chromatography), and as the increase in seed phosphate in lpa mutants compared with the wild type ranges up to tenfold, a simple high-throughput screen that tests for the ‘high inorganic P’ seed phenotype of lpa genotypes can be readily designed for use in genetics studies. This assay has subsequently been used to isolate mutants that defined a third lpa locus in maize (Shi et al., 2005b), and to isolate lpa mutations in barley (Larson et al., 1998; Rasmussen and Hatzack, 1998), rice (Larson et al., 2000), soybean (Wilcox et al., 2000) and wheat (Guttieri et al., 2004). This same highthroughput high inorganic P test also facilitates genetic mapping, and has been used to map lpa loci in maize (Raboy et al., 2000), barley (Larson et al., 1998) and rice (Larson et al., 2000; Andaya and Tai, 2005). lpa Mutations have been used to develop first-generation germplasm useful in initial evaluations of the agronomic properties and nutritional value of low-phytate types. The approach taken in these first studies was to use lpa mutations and standard ‘backcrossing’ breeding methods to develop sets of ‘near-isogenic’ lines. These sets consist of sibling lines that are either homozygous for a given wild-type non-mutant

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allele, and produce seed with normal levels of phytate and inorganic phosphate, or homozygous for the given lpa allele and produce seed with reduced phytate and increased inorganic phosphate. These sibling lines are referred to as ‘nearisogenic’ because their genomes are similar, other than allelic differences at an lpa gene. In theory, standard backcrossing methods involving a minimum of six crosses should mean that identical alleles are shared at >98% of all genes. Therefore, any differences observed in plant or seed phenotype and agronomic performance, or differences in nutritional quality, can in theory be attributed largely to the allelic difference at a single lpa gene. These types of near-isogenic lines were first produced in maize using the lpa1-1 allele of the maize lpa1 gene. Plants homozygous for lpa1-1 produce seed with a phytate reduction of about 66% compared with seed produced by wild-type non-mutant sibling lines (Table 8.1). As hybrid seed is used for maize production, near-isogenic sets of inbred lines were first produced, which were then used to produce 14 different pairs of near-isogenic hybrids (isohybrids; Ertl et al., 1998). Field trials of these first isohybrid pairs indicated that yields were reduced in eight of the lpa1-1 hybrids compared with their matching wild-type isohybrids, but yields of the lpa1-1 isohybrid in the remaining six pairs were similar to the wild type. Overall a yield reduction of about 6% was observed in the lpa1-1 hybrids, although

major differences in other aspects of plant growth and seed function were not observed. Pairs of sibling near-isogenic lines were developed using four barley lpa mutations: lpa1-1 (formerly known as M 422), lpa2-1 (formerly M 1070), lpa3-1 (formerly M 635) and M 955. Homozygosity for these mutations results in seed phytate reductions ranging from moderate (35–50%, lpa1-1 and lpa2-1) to relatively large (70–80%, lpa3-1) and extreme (>90%, M 955) (Table 8.1). Yield of isolines representing wildtype and mutant siblings were evaluated in field locations in Idaho, USA, during 2 years of regional drought (Bregitzer and Raboy, 2006; Table 8.2). These production environments were either irrigated, which resulted in relatively low drought stress, or non-irrigated (‘dryland’ or ‘rain-fed’), which resulted in considerable drought stress. The yield of isolines representing lpa1-1, lpa2-1 and lpa3-1 were statistically similar to wild-type controls when grown with irrigation. However, the yield of isolines representing the ‘extreme reduction’ mutant M 955 was clearly reduced, even in the relatively stress-free, irrigated production environments. Yield reductions associated with lpa mutations were more pronounced in the more stressful non-irrigated environments. In these environments only the yield of barley lpa1-1 was statistically indistinguishable from its wild-type sibling lines. The yield of isolines representing lpa2-1, lpa3-1 and M 955 were

Table 8.2. Comparison of yield of barley wild-type and low phytic acid sibling isolines when grown in non-stressful (irrigated) and stressful (dryland) environments.a Yield (kg/ha)c Sibling pair

Genotypeb

Seed phytic acid reduction vs. wild-type(%)

1 1 2 2 3 3 4 4

Wild-type lpa1-1 Wild-type lpa2-1 Wild-type lpa3-1 Wild-type M 955

− 50 % − 33 % − 70 % − 90 %

a

Irrigated

Dryland

8320 8487 8429 8271 7994 7991 8253 7147

1935 1718 1728 1321 1836 1353 1769 1162

A A A A A A A B

A A A B A B A B

Irrigated environments were fields at Aberdeen, Idaho, in 2002 and 2003, and Filer, Idaho, in 2003. Dryland environments were fields at Tetonia, Idaho, in 2002 and 2003, and Soda Springs, Idaho in 2003. Two replicates of six lines representing each genotype were grown at each location in a randomized complete block. b All barley lines were in the cultivar Harrington genetic background. c Values followed by the same letter within each sibling pair are not significantly different (P = 0.05).

Seed Phosphorus and Low-Phytate Crops

all substantially reduced, compared with wildtype sibling lines, when grown under drought stress. Importantly, the reductions appeared proportional to the reduction in seed phytate. These results clearly indicate that lpa mutations can be associated with yield losses and decreased stress tolerance, but that such losses are variable and are probably both functionand/or gene/allele-specific. That is, perhaps mutations in different types of functions might have a much smaller impact on yield or stress tolerance than those in other functions, genes or alleles of a given gene. Such mutations could include those in myo-inositol phosphate synthesis vs. mutations in substrate or phosphorus transport, or those giving rise to different alleles of a given gene. Barley lpa1-1 represents one case in which carefully conducted and replicated field trials indicate yields comparable to non-mutant sibling lines, possibly even in relatively stressful environments. Interestingly, the barley lpa1-1 mutation affects phytate accumulation only in the barley aleurone layer – the outer layer of the cereal grain endosperm (Ockenden et al., 2004). In cereal grains phytate accumulates in germ tissue (including the embryo and scutellum) and the aleurone, but not in the central endosperm. In seeds homozygous for barley lpa1-1, only aleurone phytate is reduced; germ phytic acid is similar to the wild type. In contrast, in seeds homozygous for barley lpa2-1, lpa3-1 and M 955, both germ and aleurone phytate are similarly reduced. Therefore, the relatively good field performance of barley lpa1-1 might be due to the fact that this mutation is in a gene of limited tissue specificity, thus reducing the impact on plant growth and performance. Alternatively, lpa1-1 might be a mutation in a transport function as opposed to a mutation in the myo-inositol phosphate pathways, and thus have a more limited impact on functions important to signal transduction and stress response, which in turn are important to yield. lpa Mutations directly impact on three cellular pools important to numerous pathways: phosphorus, myo-inositol and myo-inositol phosphates. Greatly altering these important metabolic pools could lead to yield loss. The reduced yields of lpa variants of maize and barley, compared with wild-type controls, could be due in part to a negative impact of high levels of seed inorganic phosphate on starch synthesis and accumulation.

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Starch represents the major component (>50%) of the dry weight in cereal grains, so any factor impacting on seed starch accumulation would also impact on yield. For example, a rate-limiting step in seed starch synthesis is catalysed by the enzyme adenosine diphosphate–glucose pyrophosphorylase (AGP; E.C. 2.7.7.27), which is allosterically inhibited by a high inorganic phosphate concentration (Hannah, 1997). A genetically engineered version of this enzyme that is insensitive to inorganic phosphate increased seed yield and plant biomass in rice (Smidansky et al., 2003). A useful approach to developing high-yielding lpa cultivars or hybrids might therefore be to engineer these types to express ‘deregulated’ AGP in seeds. In addition to metabolic impacts, these mutations probably have numerous downstream effects on functional properties such as stress tolerance (discussed above). Simply isolating mutations or alleles that only impart the low-phytate trait represents a first step. In addition to possible approaches that involve genetic engineering, developing high-yielding, stress-tolerant lowphytate crops will probably require classical breeding techniques, such as recurrent selection for yield and performance within lines that are homozygous lpa. This would select for combinations of ‘favourable’ alleles at numerous loci that modify or reduce the impact of the lpa genotype. This process has begun with soybeans. A soybean low-phytate mutant termed M 153 was isolated (Wilcox et al., 2000; Table 8.1) and subsequently shown to require the inheritance of recessive alleles at two non-linked loci, termed pha1 and pha2 (Oltmans et al., 2004). Crosses to several soybean lines resulted in the development of three populations (Oltmans et al., 2005). Ten wild-type lines (homozygous for dominant wild-type alleles) and ten low-phytate lines (homozygous for both pha1 and pha2 alleles) were isolated within each population. Analysis of seed traits and field performance found relatively little statistically significant differences between normal and low-phytate lines, except that the latter displayed reduced field emergence compared with sibling wild-type lines (45% vs. 68%). However, variation in field emergence within low-phytate lines indicates that positive selection may yield low-phytate soybean germplasm with acceptable field emergence. In many cases analyses of seed phosphorus fractions in lpa genotypes indicate that reductions in phytate are largely matched by increases in

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inorganic phosphate or by increases in both inorganic phosphate and other lower-order myo-inositol phosphates. This means that cellular phosphorus, defined as all forms of phosphorus other than myo-inositol phosphates and inorganic phosphate, tends to remain constant (Table 8.1). However, in some cases, such as the soybean mutant M 153 (Table 8.1), the reduction in phytate phosphorus is not matched entirely by an increase in inorganic phosphate and other myo-inositol phosphates (Oltmans et al., 2005), meaning that cellular phosphorus is increased compared with the wild type. Further analyses are required to identify the phosphorus compound(s) involved. A second study (Meis et al., 2003) revealed an intriguing downstream effect of a second type of low-phytate soybean. Hitz et al. (2002) isolated a mutant, LR33, subsequently shown to be a recessive mips allele in an MIPS loci in the soybean genome. Meis et al. (2003) observed a reduction in seedling emergence for low-phytate lines homozygous for the LR33 mips allele. However, reduced field emergence of LR33 lines compared with the wild type was influenced by seed production environment: LR33 seed produced in temperate locations such as Iowa, USA, had 63% field emergence, while seed from the same lines produced in subtropical environments such as Puerto Rico had a mean field emergence of 8%. In comparison, seed produced by wildtype sibling lines in temperate locations had a mean field emergence of 77% compared with 83% for seed produced in the subtropical environment. Homo-zygosity for the LR33 recessive mips allele therefore has an interesting and agronomically important downstream effect, because production of LR33 seed in subtropical environments results in seed with greatly reduced field emergence, even when germinated and grown in temperate locations. The cause of this effect is at present unknown.

Animal Nutrition Studies Evaluating Low-Phytate Crops In terms of animal nutrition and production, the main interest in lpa crops focuses on grain phosphorus availability in diets for non-ruminant species such as poultry, swine and fish. The

primary concerns are providing sufficient phosphorus for optimal animal growth and productivity, while reducing phosphorus concentrations in animal manure. Also of interest are calcium availability and several other measures of animal health, productivity and product quality. Perhaps the best measure of phosphorus availability in feed is the difference between phosphorus intake in feed and phosphorus excretion in manure. Phosphorus availability is also often measured indirectly in animals fed diets prepared with normal vs. low-phytate grains, by comparing various measures of animal growth rate, feed/gain ratios, bone ash and strength, and concentrations of phosphorus or calcium in bones or blood. The first published study (Ertl et al., 1998) addressing these questions compared the nutritional value of grain produced by two maize isohybrids in diets for broiler chicks. This involved grain from a ‘normal phytate’ wild-type isohybrid containing 3.8 mg total P/g dry weight (3.2 mg P/g as phytate) and a matching lpa1-1 isohybrid containing 3.9 mg total P/g dry weight (1.3 mg P/g as phytate). If we hypothesize that phytate is largely ‘non-available phosphorus’ to nonruminants, and all other forms of seed phosphorus (referred to as non-phytate phosphorus) represent available phosphorus, then the wild-type grain contained about 0.6 mg P/g as available phosphorus or 16% of grain total phosphorus, and the lpa1-1 grain contained 2.6 mg P/g as available phosphorus or 67% of grain total phosphorus. The maize wild-type and lpa1-1 experimental diets utilized in the Ertl et al. (1998) chick feeding trial were formulated identically, so the only difference between the two diets was in their phosphorus chemistry. Differences in responses to diets were therefore attributable to differential availability of phosphorus. Two diets were prepared for each of the two genotypes, wherein maize grain represented either 56% or 69% of the total diet. The additional, ‘basal’ components of the diets (soymeal, vitamins, minerals, antibiotics, etc.) contributed to about 25% of the total phosphorus, and no additional supplemental phosphorus was included. The maize component therefore represented the major source of phosphorus in the diets. All diets were otherwise formulated to contain similar, National Research Council-recommended levels (<5% difference) of other, non-phosphorus constituents providing similar energy, essential amino acids and

Seed Phosphorus and Low-Phytate Crops

minerals, including calcium (National Research Council, 1994). Estimates of available phosphorus in grain produced by the wild-type isohybrid ranged between 30% and 48%, or about 1.1–1.8 mg P/g (Ertl et al., 1998). The corresponding estimates of available phosphorus for the maize lpa1-1 grain were between 70% and 91%, or about 2.7–3.5 mg P/g. These differences parallel those in nonphytate phosphorus obtained by chemical analyses of the grains, confirming that non-phytate phosphorus is largely available to non-ruminants. As the total phosphorus concentration in grain is often similar in normal and low-phytate varieties, and phytate phosphorus normally represents a substantial fraction of grain total phosphorus, low-phytate genotypes have greatly increased available phosphorus. Optimal chick growth under the experimental conditions used in the Ertl et al. (1998) study was obtained with experimental diets prepared using monosodium phosphate as the phosphorus source to provide 0.4–0.5 mg P/g dry feed, reflecting current recommendations (National Research Council, 1994). Therefore, the diets prepared with wild-type maize provided only 20–40% of the phosphorus required for optimal growth, and this limited bird growth (bird weights at 18 days were 64–71% of those obtained from diets containing 100% recommended phosphorus). The diets formulated with lpa1-1 grain provided between 50% and 70% of the optimal level of available phosphorus, still limiting to optimal growth but less so than wild-type grain (bird weights at 18 days were 76–84% of those obtained from diets containing 100% recommended phosphorus). The objective in engineering optimized nutrient chemistry for animal agricultural production is to maximize performance or production while minimizing waste. Faecal phosphorus from birds consuming lpa1-1 maize diets was reduced by between 9% and 40% compared with that from birds consuming wild-type maize diets, and by between 30% and 47% compared with that from birds fed diets containing phosphate supplements (Ertl et al., 1998). Phosphorus nutritional status has an important impact on calcium status and bone health. It is significant that blood phosphorus and calcium levels in birds consuming the lpa1-1 maize diets were 46% and 49% greater, respectively, than in birds consuming the wild-type maize diets (Ertl

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et al., 1998). This resulted in increases of 12%, 10% and 13% in bone (tibia) ash, bone phosphorus and bone calcium, respectively. The observed differences in the availability of phosphorus and calcium and the health and productivity of the animals can be attributed to the major differences in grain phosphorus chemistry between the wild-type and lpa1-1 grain, in turn attributable in large part to a single allelic difference in the two maize genotypes. This first animal feeding trial of a low-phytate maize grain was followed by a number of studies that either confirmed or expanded upon the initial results described above: available phosphorus is increased in low-phytate grains in proportion to the increase in non-phytate phosphorus; depending on dietary formulation, use of low-phytate grains reduces animal waste phosphorus in proportion to reduced dietary phytate, with faecal phosphorus reductions between 10% and 50%; and reduced dietary phytate can enhance availability of calcium, iron and zinc. These studies used a variety of experimental approaches, including diets formulated with either low-phytate maize or low-phytate barleys, diets that evaluated products made from these grains such as low-phytate maize gluten, diets formulated using combinations of low-phytate grains and phytase enzyme supplements, and several animal systems including fish (trout), swine and poultry (chicken and turkey) (Sugiura et al., 1999; Douglas et al., 2000; Li et al., 2000, 2001; Spencer et al., 2000b; Waldroup et al., 2000; Veum et al., 2001, 2002; Peter and Baker, 2002; Jang et al., 2003; Overturf et al., 2003; Thacker et al., 2003; Yan et al., 2003). A recent development in this field was the evaluation (Sands et al., 2003; Adeola, 2005; KarrLilienthal et al., 2005) of the nutritional value of soymeal made from the M 153 low-phytate soybean described in Wilcox et al. (2000). Most of the phosphorus and phytate in the seed are concentrated in soymeal when it is processed from whole soybeans (V. Raboy, 2006, unpublished data). Phosphorus availability is increased in low-phytate soymeal compared with normal phytate soymeal, in proportion to the increase in non-phytate phosphorus (Sands et al., 2003). These studies also found that ‘metabolizable energy’ and amino acid digestibility are also increased in low-phytate soymeal compared with normal soymeal (Adeola, 2005; Karr-Lilienthal et al., 2005).

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The first published trout feeding trial (Sugiura et al., 1999) was also the first to evaluate, side by side in the same study, diets formulated with low-phytate maize (two lpa1-1 isohybrids with phytate reductions of about 66%) or lowphytate barley (an isoline of the cultivar Harrington homozygous for the barley lpa1-1 allele, formerly referred to as M 422, which results in about a 40% reduction in grain phytate; Table 8.1). This study demonstrated that when diets were formulated to contain relatively high or luxury levels of available phosphorus, most from non-grain components such as fishmeal, little benefit in terms of phosphorus or mineral availability, or reduction in waste phosphorus, was observed with the substitution of low-phytate grains for normal grains. Excess non-grain-derived dietary phosphorus overwhelmed any differences in grain-derived dietary phytate. However, when diets were formulated to maintain animal growth but minimize total dietary phosphorus, such as when using a ‘low-ash’ diet that relies more on the phosphorus provided by the grain components, benefits in phosphorus and mineral availability, including calcium, zinc and iron availability, were observed. Trout consuming these low-ash diets prepared using low-phytate grains excreted between 32% and 44% less phosphorus than did trout consuming low-ash diets prepared using ‘normal phytate’ controls. These results indicate potential use of low-phytate grains in ongoing efforts to reduce the dependence on fishmeal in aquaculture production, and to reduce aquaculture effluent phosphorus. A follow-up study with trout (Overturf et al., 2003) evaluated mineral availability in experimental diets containing barley grain (30% of total diet) produced either by the cultivar Harrington (containing ‘normal’ phytate levels of 2.45 mg P/g) or by three of the four Harrington lpa isolines listed in Table 8.1. Seed batches used for this study were produced separately from those analysed in Table 8.1 and were separately analysed for seed phosphorus and other seed constituents. Phytate contents were: barley lpa1-1 (M 422), 1.15 mg P/g (a phytate reduction of about 50%); barley lpa3-1 (M 635), 0.5 mg P/g (a phytate reduction of about 80%); barley M 955, <0.1 mg P/g (a phytate reduction of >90%). This represented the first animal or human nutrition study that evaluated more than one lpa isoline or genotype of a species in a single study. As seed total phosphorus remained fairly constant in the

four lines, the stepwise decrease in seed phytate was matched by the stepwise increase in seed non-phytate phosphorus, with other seed constituents remaining fairly constant. Sets of isolines like the barley set listed in Table 8.1 provide an experimental model with the potential to provide a precise measure of the effects of varying levels of dietary phytate or available phosphorus. In fact, Overturf et al. (2003) observed a remarkably linear increase in the apparent digestibility of dietary calcium in fish diets prepared with barley grains that occurred parallel to the predicted linear increase in apparent digestibility of dietary phosphorus and linear decreases in grain phytate. In diets prepared with Harrington (no phytate reduction), M 422 (50% reduction), M 635 (80% reduction) and M 955 (>90% reduction), the apparent digestibility of calcium was 12.9%, 27.7%, 46.6% and 59.8%, respectively. This confirms the initial finding of Ertl et al. (1998), and the results of other animal (Veum et al., 2001) and human nutrition studies (Hambidge et al., 2005), which indicated a linear, negative relationship between dietary phytate and calcium nutrition. Spencer et al. (2000a) demonstrated that growth performance of pigs fed diets consisting of lpa1-1 maize and no supplementary phosphate was equal to that observed for pigs fed normal phytate maize with supplementary phosphate. Further, when pigs were raised in a commercial facility, use of lpa1-1 low-phytate grain as a substitute for normal maize resulted in carcasses with less backfat and a higher percentage of ‘lean’ meat. This was the first indication that use of low-phytate types might enhance product quality, in addition to enhancing the management of phosphorus in animal production. A US patent was awarded subsequently, describing how the use of low-phytate maize as a feed substitute for normal maize reduces cholesterol in eggs (Stilborn et al., 2002). The nutritional mechanism leading to this unpredicted benefit of consumption of a low-phytate feed, reduced fat and cholesterol, is at present unknown.

Human Nutrition Studies Evaluating Low-Phytate Crops The greatest interest in seed-derived dietary phytate in terms of human nutrition is related to its

Seed Phosphorus and Low-Phytate Crops

impact on mineral nutrition in populations that rely on grains and legumes as staple foods (Brown and Solomons, 1991). This concern is primarily with iron and zinc, although dietary phytate can also have a negative impact on magnesium (Hurrell, 2003; Bohn et al., 2004) and calcium (Hambidge et al., 2005) nutrition. As discussed earlier, dietary phytate may also have positive roles as an anti-cancer agent, anti-oxidant and inhibitor of renal stone formation (Graf et al., 1987; Shamsuddin et al., 1988; Singh and Agarwal, 2005; Somasundar et al., 2005). However, human nutrition studies conducted to date with lpa crops have solely addressed the mineral nutrition question. The first human nutrition study conducted with a low-phytate crop evaluated iron absorption from tortillas prepared from wild-type vs. lpa1-1 maize (Mendoza et al., 1998). In this clinical-scale study, 13 non-anaemic males consumed tortillas labelled with one of two stable isotopes of iron. Apparent iron absorption was 49% greater (8.2% of intake) with tortillas prepared from lpa1-1 maize compared with the wild-type control (5.5% of intake). This result is also interesting as it pertains to the level of dietary phytate reduction necessary to observe mineral nutritional benefits. A study with soy protein (Hurrell et al., 1992) indicated that dietary phytate had to be reduced by 90% or more to observe a benefit in iron absorption. The reduction in phytate in lpa1-1 maize compared with the wild type is about twothirds, yet an improvement in iron absorption from test meals was observed by Mendoza et al. (1998). The issue of ‘threshold’ levels of reduction in dietary phytate necessary to observe nutritional benefits, and the contribution of studies utilizing lpa crops to this issue, are discussed further below. When a similar experiment was conducted with wild-type and lpa1-1 tortillas fortified with a higher level of iron (Mendoza et al., 2001), differences in iron absorption reflecting any benefit from reduced dietary phytate content were not observed. This different outcome has been interpreted as representing ‘conflicting results’ (Drakakaki et al., 2005). However, while a number of factors may have contributed to this lack of benefit observed in the Mendoza et al. (2001) follow-up study compared with the original study, such as differences in nutritional status of the test subjects, differences in the diet ingredients (higher tannins that are known to have a

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negative effect on iron retention) or methods of preparation, the most likely factor was that a much higher level of iron was provided in the test meals in the later study (4.4 mg/portion vs. 0.93 mg/portion). Therefore, rather than representing conflicting results, perhaps the difference between the results of Mendoza et al. (1998) and Mendoza et al. (2001) simply reflects something observed in the animal-model studies discussed above. Just as supplementation with phosphorus can ameliorate low phosphorus availability in animal feeds prepared with ‘normal phytate’ grains and legumes, supplementation or fortification with iron can ameliorate the negative impact of phytate in human diets. Supplementation represents an effective and established approach to provision of dietary needs, and can overcome negative impacts of dietary phytate. However, there are potential advantages to the lpa approach. This crop improvement approach uses genetics and breeding to permanently correct the problem at its source. When lpa crops are used in foods or feeds, a major need for supplementation is removed. Also, consumption of lpa feeds or foods can result in multiple or ‘global’ benefits in both phosphorus and mineral nutrition, such as enhanced iron, zinc, calcium and magnesium nutrition. In contrast, supplementation with a single mineral such as phosphorus or iron will not correct the negative effect of dietary phytate on zinc, calcium or magnesium nutrition (see below). A rapid ‘in vitro digestion/Caco-2 cell culture assay’ system was developed for testing the relative bioavailability of iron in food and meals (Yun et al., 2004). For example, dietary ascorbic acid enhances iron absorption, whereas polyphenolic compounds such as tannins inhibit iron absorption. The Caco-2 model system accurately reproduced these alternative effects on iron absorption in humans (Yun et al., 2004). Figure 8.4 illustrates the results of the in vitro digestion/Caco-2 assay when used to evaluate iron availability in wild-type and lpa1-1 maize (R.P. Glahn and V. Raboy, 2006, unpublished data). This assay is specifically for the amount of ferritin, an ironbinding protein complex that accumulates in the Caco-2 cells, shown to be an accurate measure of available iron. Available iron in wild-type maize was clearly enhanced by the inclusion of ascorbic acid (compared with cellular ferritin background levels; Fig. 8.4). However, lpa1-1 maize had available iron levels equivalent to wild-type maize plus ascorbic acid, and the addition of ascorbic acid to

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Ferritin (ng) / cell protein (mg)

40

30

20

10 Cell baseline WT

WT + Ascorbic acid

Ipa1-1 Ipa1-1 + Ascorbic acid

Fig. 8.4. Caco-2 cell culture assay of relative iron availability in grains produced by wild-type and low phytic acid (lpa) 1-1 near-isogenic maize hybrids. Relative iron availability was assayed by measurement of cellular ferritin formation. Nearisogenic hybrids were produced using wild-type (WT) and lpa1-1 versions of the inbreds A619 and A632. Phytate represented 3.1 mg P/g dry weight in wild-type (WT) grain and 1.3 mg P/g dry weight in low phytic acid (lpa) 1-1 grain. Grains were processed as described (Glahn et al., 2002; Oikeh et al., 2003) and analyses indicated that processing did not significantly impact grain phytate levels. Genotypes were tested with and without ascorbic acid (200 fM). Values represent means (n = 5) ± standard errors (P = 0.05). (Unpublished data kindly provided by Raymond P. Glahn, US Department of Agriculture, Ithaca, New York, USA.)

lpa1-1 maize further enhanced relative iron availability in a symmetric fashion. The enhanced relative iron availability of lpa1-1 maize compared with wild-type maize, as assayed by the Caco-2 model (Fig. 8.4), parallels the benefit in terms of iron availability of lpa1-1 maize observed by Mendoza et al. (1998) in a study with human subjects, further validating the Caco-2 assay (R.P. Glahn, USDA–ARS, Ithaca, New York, USA, 2006, personal communication). Further, the enhanced iron availability of lpa1-1 maize compared with its wild-type control is equivalent to, or greater than, iron bioavailability of any of 15 rice genotypes screened with the

Caco-2 assay (Glahn et al., 2002), 20 early-maturing tropical maize varieties screened with this assay (Oikeh et al., 2003) or of grains produced by maize genetically engineered to co-express soybean ferritin and Aspergillus phytase in the maize endosperm (Drakakaki et al., 2005). Thus the type of single-gene allelic change in maize lpa1-1 appears to have potential for improving the iron bioavailability of this staple food. The set of four barley lpa isolines (Table 8.1) were also evaluated in the same study (Fig. 8.4), although no difference in relative iron bioavailability was observed between wild-type and lpa sibling lines. This lack of difference among barley isolines might be due to high levels of seed tannins compared with maize, an explanation given for the difference in results between the two Mendoza et al. (1998, 2001) human nutrition studies, or to some other unknown difference in barley vs. maize grain chemistry. The first human nutrition study that used lpa maize and addressed the impact of dietary phytate on iron (Mendoza et al., 1998) was followed by several similarly designed clinical studies using small (10–20) numbers of volunteer subjects that addressed the impact on zinc and calcium. In the case of zinc, two independently developed lpa maize genotypes were compared with their respective wild-type isohybrids as controls: maize lpa1-1 grain with a 60% reduction in phytate; and ‘NutriDense Low-Phytate’, a maize hybrid producing grain with an 80% reduction in phytate (Adams et al., 2002; Hambidge et al., 2004). Previous studies of the impact of dietary phytate on zinc nutrition indicated that the molar ratio of dietary phytate to zinc is critical to zinc availability. This prior work defined a 10:1 dietary phytate/zinc molar ratio threshold above which negative impacts of phytate on zinc retention were predicted. The phytate/zinc molar ratios measured for the grain produced by the four genotypes used in these studies were 37:1 (wild-type lpa1-1 control), 28:1 (wild-type NutriDense Low-Phytate control), 17:1 (lpa1-1) and 7:1 (NutriDense Low-Phytate) (Hambidge et al., 2004). The corresponding values for ‘fractional zinc absorption’, measured following consumption, by ten volunteers, of test meals prepared with different stable isotopes of zinc as markers for a given genotype, were 0.151 ± 0.071 (wild-type lpa1-1 control), 0.135 ± 0.050 (wild-type NutriDense Low-Phytate control),

Seed Phosphorus and Low-Phytate Crops

0.285 ± 0.042 (lpa1-1) and 0.383 ± 0.066 (NutriDense Low-Phytate). Thus, studies of both zinc and iron availability from lpa genotypes (Mendoza et al., 1998) indicated a linear relationship, rather than a critical threshold, between phytate/zinc molar ratio and fractional zinc absorption, and between dietary phytate and iron availability. This is of practical importance to nutrition science and crop improvement when addressing issues of mineral nutrition in populations dependent on grains and legumes as staple foods. For example, it indicates that even moderate heritable reductions in grain or legume seed phytate might result in valuable improvements in both iron and zinc nutrition when these crops are used as basic foods. Hambidge et al. (2005) evaluated the absorption of calcium from tortilla meals prepared using grain produced by the same maize lpa1-1 isoline, compared with its appropriate wild-type control, used in the above zinc nutrition study. Similar clinical-scale experimental methods were used, including stable calcium isotopes as tracers of lpa grain type, and the volunteer subjects were five healthy adult women. Mean fractional calcium absorptions for wild-type or lpa1-1 maize tortillas were 0.35 ± 0.07 and 0.50 ± 0.03, respectively, a statistically significant difference (P = 0.003). These clinical-scale and model system studies provide important first-generation data on the potential value, in terms of nutritional enhancement, of genetic changes in seed crops, whether accomplished through classical genetics or genetic engineering. However, these small-scale (in terms of numbers of participants), short-duration studies must be followed by larger-scale (more participants), longer-duration field studies, in which participants consume diets prepared with wild-type and lpa seeds in a non-clinical setting. The smallscale, short-duration clinical studies measure acute phenomena, whereas the larger-scale longer-duration field studies would measure impacts on health and nutrition of chronic consumption of diets differing in phytate levels. The first such study (Mazariegos et al., 2006) found no clear and large difference in zinc absorption, or any statistically significant difference, in healthy Guatemalan children (whose traditional diets rely on maize as a staple food), when consuming either wild-type or lpa1-1 maize for a 10-week period. However, children consuming the lpa1-1 maize compared with children consuming the

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wild-type control did display higher fractional zinc absorption (14%). The difference was not statistically significant, indicating that the differences in dietary phytate in this context had no great impact on zinc absorption. It could also reflect other factors, such as the fact that differences in dietary phytate consumed between the test groups were less than expected due to other sources of phytate in the diet, smaller than expected differences in dietary phytate provided by the different maize types and higher than expected levels of zinc absorption by test subjects. This highlights the challenge represented by the need to conduct longer-duration, larger-scale studies. It is possible that the results of small-scale clinical studies that indicate a benefit to reduced dietary phytate do not translate into real-world benefits or, alternatively, that it is inherently more challenging to accurately measure differences in nutritional status in larger-scale field studies. Whatever the explanation for these differences, it is ultimately essential to prove a benefit under real-world conditions. Studies addressing the impact of reductions in grain or legume phytate in lpa versions of various crops on magnesium nutrition in humans have not yet been conducted. However, a recent finding concerning the distribution of minerals in white rice prepared from lpa1-1 rice compared with white rice prepared from the wild-type control has relevance to magnesium nutrition. In cereal grains such as rice and wheat, most phytate and minerals, including potassium and magnesium, are deposited in the germ and outer aleurone layers of the seed, mostly as mixed phytate salts. These outer layers are removed during milling as the bran fraction. The central endosperm, which following whole grain milling ends up as white rice and white wheat flower, contains very low levels of phosphorus and minerals. It is possible that a mutation that blocks the ability of seeds to make phytate might alter the distribution of minerals in the mature seed. In fact, Bryant et al. (2005) found that phosphorus, potassium and magnesium levels in milled white rice prepared from lpa1-1 rice, a mutation that reduces seed phytate by about 40% (Table 8.1), were increased from 25% to 40% compared with levels in milled white rice prepared from the wild-type control. Thus, an additional benefit to the lpa approach may prove to be that lpa mutations can favourably alter the distribution of

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minerals in grains such as wheat and rice that are milled prior to consumption. In contrast to the results of Bryant et al. (2005), an analysis of fractions obtained following milling of the wheat lpa mutant Js-12-LPA found no large differences in phosphorus and mineral contents between the mutant and the wild-type control (Guttieri et al., 2004). Clearly, more work is needed to evaluate the impact of lpa mutations on grain mineral content and distribution. Both of the above studies evaluated only one mutant, and in each case this mutant only conditioned a ‘moderate’ block (<50%) in grain phytate. It would be interesting to conduct a similar analysis of milling fractions obtained from the set of barley lpa mutants listed in Table 8.1 that condition blocks in grain phytate accumulation of up to 90%. Milling analysis of a mutant such as barley M 955, in which grain phytate is nearly absent, should provide a definitive test of the hypothesis that blocking grain phytate synthesis can greatly alter grain mineral distribution. Three additional studies (Liu et al., 2004; Ockenden et al., 2004; Joyce et al., 2005) reported the distribution of phosphorus and minerals in grain fractions from the rice, wheat and barley lpa mutants listed in Table 8.1, and found little difference between wild-type and lpa fractions. However, these studies took a different approach to that of Bryant et al. (2005) and Guttieri et al. (2004) in that they analysed fractions obtained following dissection of grains into two fractions: ‘germ’, which included the embryo and scutellum; and ‘rest-of-grain’, which included both the central endosperm and aleurone layer. Therefore, these later studies could not detect any differences in mineral distribution between the aleurone layer and the endosperm, as could the studies analysing milling fractions.

Seed Total Phosphorus and Ruminant Nutrition While dietary phytate is not thought of as a major issue in ruminant nutrition, there is ongoing evaluation of the possible benefits of reduced dietary phytate, or the use of phytase as a feed supplement, in this production context. For example, one recent study indicated that phytase use as a feed supplement for ruminants

might improve phosphorus availability (Kincaid et al., 2005). As seed phosphorus chemistry is not a major issue in some ruminant production systems, a new direction might be to develop genetic resources useful for breeding crops with reduced seed total phosphorus. Dairy and beef cattle can often consume more phosphorus than needed for optimal production, which, as in non-ruminant production, leads to problems in manure management (Volk et al., 2000; Erickson et al., 2002; Rotz et al., 2002; Toor et al., 2005). Heritable reductions in seed total phosphorus from 25% to 50% might translate into reductions in dairy and beef cattle waste phosphorus of similar magnitude. Breeding efforts using traditional methods are currently underway to select for ‘reduced seed total phosphorus’ maize lines (Warden and Russell, 2004). To date, most lpa mutations have greatly altered seed phosphorus chemistry, but not greatly altered seed total phosphorus. In fact, most lpa mutations so far have been isolated by screening for the high seed inorganic phosphate phenotype, a phenotype most pronounced and therefore most often identified in a selection, when seeds have normal levels of total phosphorus but greatly reduced levels of phytate. However, one barley lpa mutation, barley lpa1-1, consistently has reduced seed total phosphorus compared with wild-type controls, typically ranging from 10% to as high as 23% (Table 8.1; Dorsch et al., 2003; Ockenden et al., 2004; V. Raboy, 2006, unpublished data). It is possible that a previously unnoticed class of lpa mutations might be those that alter phosphorus transport and impact seed total phosphorus. The US Department of Agriculture–Agricultural Research Service collection of barley lpa mutations is being re-evaluated in this light. One might also conduct a second-generation genetic screen for single-gene mutations that reduce seed total phosphorus but that do not impact plant phosphorus, such as mutations that block the transport of phosphorus from parent plant to progeny seed. One could screen progeny from mutagenized populations directly for reduced seed total phosphorus (Fig. 8.5, left, screen no. 1). As is obvious in Fig. 8.5, a ‘low total P’ seed mutant would also be ‘low-phytate’ and, as for barley lpa1-1, may represent a valuable alternative approach to the dietary problems associated with phytate. Alternatively, one could screen progeny of chemically mutagenized lpa

Seed phosphate fraction (mg P/g)

Seed Phosphorus and Low-Phytate Crops

4 Screen no. 1

Screen no. 2

2

0

Hypothetical low total P

Phytate P

Low phytic Hypothetical acid low phytic acid / low total P Genotype

Normal

Inorganic P

Other P

Fig. 8.5. Two types of screens for reduced seed total phosphorus. Seed phosphorus fractions in four types of grain genotypes: homozygous wild-type or ‘normal’; low phytic acid (lpa); a hypothetical low total P; a hypothetical low phytic acid P/low total P. The dashed arrows indicate the two types of genetic screens currently underway: selection for seeds with reduced seed total phosphorus (screen no. 1) vs. selection for reduced inorganic phosphate in low phytic acid seeds that normally have high inorganic phosphate (screen no. 2).

lines for those mutations that reduce the high inorganic phosphate phenotype of lpa seeds (Fig. 8.5, right, screen no. 2). Mutations or alleles that reduce the high inorganic phosphate of lpa seeds would represent one type of reversion of the phenotype of lpa mutants. Reduced seed inorganic phosphate might also reduce the negative impact of lpa mutations on yield, as the high inorganic phosphate levels in lpa seeds might suppress starch accumulation, which is so important to yield. This would also represent a type of reversion of the lpa phenotype. It is therefore possible that in the future genetics resources that reduce seed total phosphorus might be used in combination with lpa alleles to engineer optimal nutritional quality for seed crops.

Summary and Future Directions Substantial progress has been made in the molecular biology and genetics of seed phytate. Lowphytate types of soybean and several major grain crops are available for evaluation. Efforts at breeding high-yielding low-phytate grain with

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good agronomic characteristics and stress tolerance are underway. Future work in the development of high-yielding low-phytate crops will focus on continued breeding with currently available genetic resources, but may also involve engineering ‘optimized’ lpa crops by directing efforts at selected candidate genes (Stevenson-Paulik et al., 2005) and by using molecular genetics to target gene expression changes to specific target tissues such as the germ or aleurone layer. In this way optimal reductions in seed phytate can be achieved while minimizing the impact of changes in gene expression in non-seed tissues and other whole-plant processes. For example, when a mutation in a myo-inositol phosphate kinase blocks seed phytate synthesis, it might also negatively impact the way roots and shoots respond to stress. By targeting a block in a myo-inositol kinase activity in the seed only, the negative impact on vegetative stress response might be avoided. In the area of crop improvement, future studies will also focus on developing genetic resources useful in breeding ‘low seed total P’ crops. The most clear-cut application for such genotypes is in ruminant production systems such as dairy and beef production. However, they might also prove useful in engineering lowphytate crops in which the negative impact on starch accumulation and yield, perhaps one outcome of the high inorganic phosphate phenotype of low-phytate types, is greatly reduced. A growing number of human and animal nutrition studies evaluating lpa crops indicate that genetic reductions in crop seed phytate may have broad benefits in animal and human nutrition. This includes benefits in phosphorus management in non-ruminant livestock production, improvements in product nutritional quality and benefits in mineral nutrition in human populations that rely on cereals and legumes as staple foods. Future directions in the human nutrition field might include additional field-scale studies to evaluate the potential value of low-phytate crops in a non-clinical setting, involving much larger numbers of participants and a longer duration than typical clinical-scale studies. Each of the initial generation of clinical-scale human nutrition studies focused on the impact of dietary phytate on a single mineral nutrient. Future studies might therefore address mineral nutrition and health in broader terms, for example by evaluating iron,

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zinc, calcium and magnesium nutrition simultaneously in the same subjects. Future studies might also utilize sets of isolines that differ greatly in seed phytate to evaluate the potential positive roles of dietary phytate as an anti-oxidant and anti-cancer agent (Graf et al., 1987; Shamsuddin et al., 1988; Singh and Agarwal, 2005; Somasundar et al., 2005), or as an inhibitor of renal stone formation (Grases et al., 2000). Past studies that addressed the potential positive roles of dietary phytate have not moved far beyond in vitro assays, cell culture studies or

whole-animal models other than the rat (Vucenik and Shamsuddin, 2003). The ability to produce substantial amounts of grains or legumes that have large differences in endogenous phytate, but that are otherwise nearly identical, provides an excellent model to test the various hypotheses concerning the potential positive role of dietary phytate. For example, ready access to sufficient quantities of low-phytate grains and legumes would permit large-scale, long-duration studies that use large animals. Such studies would serve well as a model for human nutrition and health.

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Li, Y.C., Ledoux, D.R., Veum, T.L., Raboy, V. and Zyla, K. (2001) Low phytic acid barley improves performance, bone mineralization and phosphorus retention in turkey poults. Journal of Applied Poultry Research 10, 178–185. Liu, J.C., Ockenden, I., Truax, M. and Lott, J.N.A. (2004) Phytic acid-phosphorus and other nutritionally important mineral nutrient elements in grains of wild-type and low phytic acid (lpa1-1) rice. Seed Science Research 14, 109–116. Loewus, F.A. and Murthy, P.P.N. (2000) myo-Inositol metabolism in plants. Plant Science 150, 1–19. Loewus, M.W., Sasaki, K., Leavitt, A.L., Munsell, L., Sherman, W.R. and Loewus, F.A. (1982) The enantiomeric form of myo-inositol-1-phosphate produced by myo-inositol-1-phosphate synthase and myo-inositol kinase in higher plants. Plant Physiology 70, 1661–1663. Lott, J.N.A., Greenwood, J.S. and Batten, G.D. (1995) Mechanisms and regulation of mineral nutrient storage during seed development. In: Kigel, J. and Galili, G. (eds) Seed Development and Germination. Marcel Dekker, New York, pp. 215–235. Lott, J.N.A., Ockenden, I., Raboy, V. and Batten, G.D. (2000) Phytic acid and phosphorus in crop seeds and fruits: a global estimate. Seed Science Research 10, 11–33. Maeshima, M. (2000) Vacuolar H+-pyrophosphatase. Biochimica et Biophysica Acta 1465, 37–51. Mazariegos, M., Hambidge, K.M., Krebs, N.F., Westcott, J.E., Lei, S., Grunwald, G.K., Campos, R., Barahona, B., Raboy, V. and Solomons, N.W. (2006) Zinc absorption in Guatemalan schoolchildren fed normal or low-phytate maize. American Journal of Clinical Nutrition 83, 59–64. Meis, S.J., Fehr, W.R. and Schnebly, S.R. (2003) Seed source effect on field emergence of soybean lines with reduced phytate and raffinose saccharides. Crop Science 43, 1336–1339. Mendoza, C., Viteri, V.E., Lönnerdal, B., Young, K.A., Raboy, V. and Brown, K.H. (1998) Effect of genetically modified, low-phytic acid maize on absorption of iron from tortillas. American Journal of Clinical Nutrition 68, 1123–1128. Mendoza, C., Viteri, F.E., Lönnerdal, B., Raboy, V., Young, K.A. and Brown, K.H. (2001) Absorption of iron from unmodified maize and genetically altered, low-phytate maize fortified with ferrous sulfate or sodium iron EDTA. American Journal of Clinical Nutrition 73, 80–85. National Research Council (1994) Nutrient Requirements of Domestic Animals, 9th edn. National Academy Press, Washington, DC. Ockenden, I., Dorsch, J.A., Reid, M.M., Lin, L., Grant, L.K., Raboy, V. and Lott, J.N.A. (2004) Characterization of the storage of phosphorus, inositol phosphate and cations in grain tissues of four barley (Hordeum vulgare L.) low phytic acid genotypes. Plant Science 167, 1131–1142. O’Dell, B.L., de Boland, A.R. and Koirtyohann, S.R. (1972) Distribution of phytate and nutritionally important elements among the morphological components of cereal grains. Journal of Agricultural and Food Chemistry 20, 718–721. Oikeh, S.O., Menkir, A., Maziya-Dixon, B., Welch, R. and Glahn, R.P. (2003) Assessment of concentrations of iron and zinc and bioavailable iron in grains of early-maturing tropical maize varieties. Journal of Agricultural and Food Chemistry 51, 3688–3694. Oltmans, S.E., Fehr, W.R., Welke, G.A. and Cianzio, S.R. (2004) Inheritance of low-phytate phosphorus in soybean. Crop Science 44, 433–435. Oltmans, S.E., Fehr, W.R., Welke, G.A., Raboy, V. and Peterson, K.L. (2005) Agronomic and seed traits of soybean lines with low-phytate phosphorus. Crop Science 45, 593–598. Overturf, K., Raboy, V., Cheng, Z. J. and Hardy, R.W. (2003) Mineral availability from barley low phytic acid grains in rainbow trout (Oncorhynchus mykiss) diets. Aquaculture Nutrition 9, 239–246. Peter, C.M. and Baker, D.H. (2002) Bioavailability of phosphorus in corn gluten feed derived from conventional and low-phytate maize. Animal Feed Science and Technology 95, 63–71. Phillippy, B.Q., Ullah, A.H. J. and Ehrlich, K.C. (1994) Purification and some properties of inositol 1,3,4,5,6pentakisphosphate 2-kinase from immature soybean seeds. Journal of Biological Chemistry 269, 28393–28399. Raboy, V. (1997) Accumulation and storage of phosphate and minerals. In: Larkins, B.A. and Vasil, I.K. (eds) Cellular and Molecular Biology of Plant Seed Development. Kluwer Academic Publishers, Dordrecht, The Netherlands, pp. 441–477. Raboy, V. (2003) myo-Inositol-1,2,3,4,5,6-hexakisphosphate. Phytochemistry 64, 1033–1043. Raboy, V., Gerbasi, P.F., Young, K.A., Stoneberg, S.D., Pickett, S.G., Bauman, A.T., Murthy, P.P.N., Sheridan, W.F. and Ertl, D.S. (2000) Origin and seed phenotype of maize low phytic acid 1-1 and low phytic acid 2-1. Plant Physiology 124, 355–368. Rasmussen, S.K. and Hatzack, F. (1998) Identification of two low-phytate barley (Hordeum vulgare L.) grain mutants by TLC and genetic analysis. Hereditas 129, 107–112.

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Rotz, C.A., Sharpley, A.N., Satter, L.D., Gburek, W.J. and Sanderson, M.A. (2002) Production and feeding strategies for phosphorus management on dairy farms. Journal of Dairy Science 85, 3142–3153. Sands, J.S., Ragland, D., Wilcox, J.R. and Adeola, O. (2003) Relative bioavailability of phosphorus in low-phytate soybean meal for broiler chicks. Canadian Journal of Animal Science 83, 95–100. Shamsuddin, A.M., Elsayed, A.M. and Ullah, A. (1988) Suppression of large intestinal cancer in F344 rats by inositol hexaphosphate. Carcinogenesis 9, 577–580. Sharpley, A.N., Chapra, S.C., Wedepohl, R., Sims, J.T., Daniel, T.C. and Reddy, K.R. (1994) Managing agricultural phosphorus for protection of surface waters: issues and options. Journal of Environmental Quality 23, 437–451. Shears, S.B. (2004) How versatile are inositol phosphate kinases? Biochemical Journal 377, 265–280. Shi, J., Wang, H., Wu, Y., Hazebroek, J., Meeley, R.B. and Ertl, D.S. 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(1993) The detection, purification, structural characterization and metabolism of diphosphoinositol pentakisphosphate(s) and bisdiphosphoinositol tetrakisphosphate(s). Journal of Biological Chemistry 268, 4009–4015. Stevenson-Paulik, J., Odom, A.R. and York, J.D. (2002) Molecular and biochemical characterization of two plant inositol polyphosphate 6-/3-/5-kinases. Journal of Biological Chemistry 277, 42711–42718. Stevenson-Paulik, J., Bastidas, R.J., Chiou, S.-T., Frye, R.A. and York, J.D. (2005) Generation of phytate-free seeds in Arabidopsis through disruption of inositol polyphosphate kinases. Proceedings of the National Academy of Science of the United States of America 102, 12612–12617. Stilborn, H.L, Crum, R.C., Rice, D.W., Saunders, C.A., Hinds, M.A., Ertl, D.S., Beach, L.R., Huff, W.E. and Kleese, R.A. (2002) Method of reducing cholesterol in eggs. US Patent No. 6,391,348 B1. Sugiura, S.H., Raboy, V., Young, K.A., Dong, F.M. and Hardy, R.W. 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Vucenik, I. and Shamsuddin, A.M. (2003) Cancer inhibition by inositol hexaphosphate (IP6) and inositol: from laboratory to clinic. Journal of Nutrition 133, 3778S–3784S. Waldroup, P.W., Kersey, J.H., Saleh, E.A., Fritts, C.A., Yan, F., Stilborn, H.L., Crum, R.C. Jr and Raboy, V. (2000) Nonphytate phosphorus requirement and phosphorus excretion of broiler chicks fed diets composed of normal or high available phosphate corn with and without microbial phytase. Poultry Science 79, 1451–1459. Warden, B.M. and Russell, W.K. (2004) Resource allocation in a breeding program for phosphorus concentration in maize grain. Crop Science 44, 753–757. Wilcox, J., Premachandra, G., Young, K. and Raboy, V. (2000) Isolation of high seed inorganic P, low phytic acid soybean mutants. Crop Science 40, 1601–1605. Wilson, M.P. and Majerus, P.W. (1996) Isolation of inositol 1,3,4-trisphosphate 5/6-kinase, cDNA cloning and expression of the recombinant enzyme. Journal of Biological Chemistry 271, 11904–11910. Wilson, M.P. and Majerus, P.W. (1997) Characterization of a cDNA encoding Arabidopsis thaliana inositol 1,3,4trisphosphate 5/6-kinase. Biochemical and Biophysical Research Communications 232, 678–681. Yan, F., Fritts, C.A., Waldroup, P.W., Stilborn, H.L., Rice, D., Crum, R.C. Jr and Raboy, V. (2003) Comparison of normal and high available phosphorus corn with and without phytase supplementation in diets for male large white turkeys grown to market weights. International Journal of Poultry Science 2, 83–90. York, J.D., Odom, A.R., Murphy, R., Ives, E.B. and Wente, S.R. (1999) A phospholipase C-dependent inositol polyphosphate kinase pathway required for efficient messenger RNA export. Science 285, 96–100. Yoshida, K.T., Wada, T., Koyama, H., Mizobuchi-Fukuoka, R. and Naito, S. (1999) Temporal and spatial patterns of accumulation of the transcript of myo-inositol-1-phosphate synthase and phytin-containing particles during seed development in rice. Plant Physiology 119, 65–72. Yun, S., Habicht, J.-P., Miller, D.D. and Glahn, R.P. (2004) An in vitro digestion/Caco-2 cell culture system accurately predicts the effects of ascorbic acid and polyphenolic compounds on iron bioavailability in humans. Journal of Nutrition 134, 2717–2721.

9

Phytase and Inositol Phosphates in Animal Nutrition: Dietary Manipulation and Phosphorus Excretion by Animals Xin Gen Lei1 and Jesus M. Porres2 1

Department of Animal Science, Morrison Hall 252, Cornell University, Ithaca, NY 14853, USA; 2Departamento de Fisiología, Universidad de Granada, Granada, Spain

Salts of myo-inositol hexakisphosphate (phytate) represent between 60% and 80% of the total phosphorus in plant seeds used to feed animals (see Raboy, Chapter 8, this volume), but simple-stomached species such as swine, poultry and fish do not have hydrolytic enzymes in their upper digestive tracts to digest phytate in feed. These species therefore require dietary supplemental inorganic phosphate to maintain productivity, but the unutilized phytate is excreted, resulting in high phosphorus concentrations in manure that exceed the phosphorus requirements of most crops ( Whalen and Chang, 2001; Adeli et al., 2005). The manure phosphorus therefore accumulates in soil and increases the risk of phosphorus pollution of water bodies (see Leytem and Maguire, Chapter 10, this volume). The impact of phosphorus pollution from animal manures is being exacerbated by the rising global demand for meat, which has resulted in animal production being consolidated into large intensive feeding operations to improve efficiency. In addition to environmental issues, myo-inositol hexakisphosphate chelates divalent metals such as calcium, zinc and iron (Cheryan, 1980), which renders these nutrients unavailable to simplestomached humans and animals. To address environmental concerns surrounding phosphorus pollution from animal operations, considerable research has been directed towards manipulating animal diets to improve the digestibility of phosphorus in grain feed and mini-

mize phosphorus excretion in manures. This chapter reviews research on manipulation using phytase, currently the most common strategy to improve phosphorus efficiency in animal production. Further details on phytase can be found elsewhere in this volume (see Greiner, Chapter 6, and Mullaney and Ullah, Chapter 7), and the development of low-phytate grains, an alternative strategy of dietary manipulation to reduce manure phosphorus, is also discussed (see Raboy, Chapter 8, this volume).

Manipulation of Phosphorus Nutrition and Excretion The commonly used feedstuffs, maize and soybean meal, contain ~0.8–1.1% and 1.3–2.2% phytate, respectively. Protein products such as soy isolate, canola, sunflower or cottonseed meal contain 1.3–5.0% phytate (Han, 1988; Eeckhout and De Paepe, 1994; Fernández-Quintela et al., 1997; Kasim and Edwards, 1998; Leske and Coon, 1999; Ravindran et al., 1999a; Shen et al., 2005). Bioavailability of phosphorus from these feedstuffs, except for those with relatively high intrinsic phytase activity (Eeckhout and De Paepe, 1994; Viveros et al., 2000; Zimmermann et al., 2002; Shen et al., 2005), is <15% for swine (Cromwell, 1992; Weremko et al., 1997) and

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15–40% for poultry (Leske and Coon, 1999). Nevertheless, the digestibility of total phosphorus or phytate for a given feed ingredient may be affected by diet composition, age of the animal, experimental protocol (i.e. faecal vs. ileal digestibility), amount of phosphorus present in the diet or endogenous excretion of phosphorus (Shen et al., 2002). Several strategies have been applied to reduce phosphorus excretion by animals and its accumulation in soil (Fig. 9.1). Meeting the exact nutrient requirements of phosphorus by different species is the first step to reducing their phosphorus excretion, and digestible phosphorus levels are more accurate than total phosphorus levels for that purpose (Poulsen et al., 1999). Phase feeding with accurate dietary phosphorus allowances is another cost-effective way to reduce faecal phosphorus excretion. High levels of available phosphorus are required during the early stages of growth and development of the animals, whereas much lower levels are needed at later stages (Keshavarz, 2000). Therefore, the amount of phosphorus added to the feed can be significantly reduced across the growing–finishing period (Applegate et al., 2003a; Angel et al., 2005). It is also important to recognize the greater efficiency of commercial herds or strains in utilizing nutrients (Havenstein et al., 2003) and their ability to adapt to a moderate deficiency of phosphorus (Yan et al., 2005). Feed supplements such as vitamin D and its analogues or citric acid can improve phosphorus utilization, thereby reducing its excretion (Boling et al., 2000; Snow et al., 2004). The recent development of low phytate or high available phosphorus crops (Spencer et al., 2000;

Accurate estimate of phosphorus requirements

Diet optimization Phase feeding

Reduction of manure phosphorus excretion

Low-phytate crops

Use of supplemental phytases

Fig. 9.1. Strategies for reducing manure phosphorus excretion.

Veum et al., 2001; see Raboy, Chapter 8, this volume) has provided hope for a simple and sustainable solution. Commercially available manure amendments like alum can significantly reduce soluble phosphorus in poultry litter (Moore et al., 1999). Best management practices have been developed to prevent diffuse pollution of surface and ground waters by agricultural phosphorus (Sharpley et al., 2001). Above all, dietary supplementation of a microbial enzyme, phytase, has proven to be the most effective tool for animal industry to reduce phosphorus excretion from animal waste to comply with the environmental regulations.

Nutritional Impacts of Phytase Phytase is a phosphohydrolytic enzyme that initiates the stepwise removal of phosphate from myoinositol hexakisphosphate (see Mullaney and Ullah, Chapter 7, this volume). Numerous experiments have shown that supplemental microbial phytase at 300–1000 units/kg in swine and poultry diets improves phosphorus bioavailability by 10–35% (Lei and Stahl, 2000). Animal response to dietary phytase dose monitored by growth performance, apparent phosphorus absorption, plasma inorganic phosphorus concentration, plasma alkaline phosphatase activity, and bone ash and breaking strength was either linear or curvilinear (Lei et al., 1993a,b; Kornegay and Qian, 1996; Yi et al., 1996; Gentile et al., 2003). The general estimate is that inclusion of 300–600 phytase units/kg in swine and poultry diets releases 0.8 g of digestible phosphorus and replaces either 1.0 or 1.3 g of phosphorus from mono- and dicalcium phosphate, respectively (Ravindran et al., 1995; Yi et al., 1996; Radcliffe and Kornegay, 1998; EsteveGarcia et al., 2005). Apparently, the inorganic phosphorus equivalence of phytase can be affected by the physiological status, housing and behaviour (coprophagy) of the animals (Kemme et al., 1997a,b), biochemical properties of the enzyme (Augspurger et al., 2003; Applegate et al., 2003b) and the nutritional indices or parameters chosen to estimate phosphorus availability and phytase efficacy (Esteve-Garcia et al., 2005). Because of the acidic pH optimum and greater pepsin resistance, bacterial phytase (AppA2) is

three- to fourfold more effective than fungal PhyA phytase in both swine and poultry diets (Augspurger et al., 2003; Applegate et al., 2003b). Supplemental phytase also improves utilization of other minerals by animals (Lei and Stahl, 2000, 2001; Debnath et al., 2005; Lei and Porres, 2005) and humans (Sandberg et al., 1996; Porres et al., 2001, 2005). The potential of phytase to improve iron availability in human foods is of special interest due to the worldwide prevalence of iron deficiency.

Phosphorus Reduction in Manure from Animals Fed Modified Diets By improving the bioavailability of dietary phytate, supplemental phytase significantly reduces phosphorus excretion by animals. The decreases range between 10% and 50%, depending on dietary phosphorus concentration, supplemental phytase activity and diet composition ( Jongbloed et al., 1992; Lei et al., 1993a,b; Yi et al., 1996; Liu et al., 1997; Baxter et al., 2003; Applegate et al., 2003b). According to Jongbloed and Lenis (1992), a pig excretes a total of 1.23 kg of phosphorus during its life cycle. If supplemental phytase produces an average of 30% reduction in manure phosphorus, ~22,000 t of manure phosphorus annually in the USA from raising marketing pigs alone (60 million pigs marketed/year in the USA × 0.37 kg phosphorus/head) would be prevented from entering the environment. Supplementation of phytase and the use of low-phytate grains both decrease the amount of total phosphorus present in manure-amended soils (Maguire et al., 2003). In addition, supplemental phytase reduces faecal excretion of calcium by up to 50% (Fig. 9.2; Lei et al., 1993a) and presumably other minerals as well (Lei et al., 1993c; Stahl et al., 1999). Nevertheless, effects of phytase and low-phytate grains on water-soluble phosphorus excretion remain unclear. Gollany et al. (2003) reported a 42% reduction in total manure phosphorus in pigs fed low-phytate maize and no differences in manure phosphorus solubility when compared with pigs fed regular maize. Similar findings were reported by Applegate et al. (2003a) in poultry fed low-phytate maize or phytase. Wienhold and Miller (2004) did not find significant differences in the distribu-

% on dry basis

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Basal + 750 units/kg

1

0 Faecal phosphorus

Faecal calcium

Fig. 9.2. Reduction of faecal phosphorus and calcium concentrations in pigs fed 750 units of phytase/kg of maize–soybean meal diet compared with those fed only the maize–soybean meal diet. (From Lei et al., 1993a.)

tion of phosphorus fractions in manure from swine fed low-phytate or regular maize, whereas Penn et al. (2004) observed considerable reductions in the amount of phosphorus runoff generated under stimulated rainfall from soils amended with manure from turkey fed phytase or lowphytate maize compared with those fed the standard diet. Baxter et al. (2003) found lower levels of total and soluble phosphorus in slurries of pigs fed phytase, low-phytate maize or a combination of both when compared with those fed a standard diet. However, the dietary treatments of phytase or low-phytate maize resulted in a higher proportion of dissolved molybdate reactive phosphorus in the slurries. The environmental effects of dietary manipulation on phosphorus solubility and transfer are discussed in detail elsewhere (see Leytem and Maguire, Chapter 10, this volume).

Functional Site of Phytase in the Animal Two research groups ( Jongbloed et al., 1992; Yi and Kornegay, 1996) showed the stomach of pigs to be the major site of supplemental Aspergillus niger PhyA phytase activity. Low activity was detected in the proximal small intestine, and negligible amounts in the distal small intestine. Our recent work with a bacterial phytase Escherichia coli AppA2 (Pagano A.R. et al., 2005, unpublished data) has illustrated that stomach is also the main functional site of the enzyme. Because of its stronger pepsin resistance compared to PhyA (Rodriguez et al., 1999a), pigs fed AppA2 maintained similar phytase activity in digesta among

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stomach, duodenum and upper jejunum. By the lower jejunum, all supplemental phytase disappeared in digesta. However, there was a significant inverse relationship between colon phytase activity and dietary phytase supplementation, indicating the strong dependence of colonial microbial phytases on phytate substrate. Similar findings were reported by Schlemmer et al. (2001). Likewise, the crop, gizzard and proventriculus are the major sites of supplemental activity in poultry, and little activity is found in the small intestine (Yu et al., 2004). The total inactivation or degradation of supplemental phytase in the small intestine of swine and poultry precludes any adverse effect of the enzyme in releasing soluble phosphorus into the environment upon storage of manure prior to spreading it into the fields. The reduced colon microbial phytase activity in animals fed dietary supplemental phytase will also decrease free or soluble phosphorus in the colon and in manure.

Determinants of Dietary Phytase Efficacy A number of factors can modulate the effectiveness of supplemental phytase in diets for swine or poultry (Fig. 9.3). The most consistent factor, and probably the strongest, is the dietary cal-

Ca/P ratio Inorganic P

Vitamin D derivatives

cium/phosphorus ratio. A ratio of 2:1 or wider has adversely affected phytase function in both swine and poultry, compared with ratios close to 1:1 (Fig. 9.4) (Lei et al., 1994; Qian et al., 1997; Liu et al., 1998; Tamim et al., 2004), which is probably due to the precipitation of calcium phytate. Therefore, a 1.2:1 ratio of calcium/phosphorus is recommended for phytase-supplemented diets (Liu et al., 1998). Likewise, adding inorganic phosphorus to phytase-supplemented diets may reduce the efficacy of the enzyme due to product inhibition. Several groups have demonstrated a positive effect of vitamin D derivatives on the utilization of total phosphorus, phytate-phosphorus, calcium, zinc and manganese by poultry (e.g. Edwards, 1993; Biehl et al., 1995). Vitamin D may act synergistically with phytase on dietary calcium and phosphorus retention in broilers (Biehl et al., 1995; Qian et al., 1997; Snow et al., 2004; Angel et al., 2005), although the effect seems to be more pronounced in poultry than in swine (Lei et al., 1994; Biehl and Baker, 1996). Supplementation of swine or poultry diets with citric, formic or lactic acid effectively improves daily gain, feed-use efficiency and apparent total tract digestibility of organic matter, ash, phosphorus, calcium and magnesium (Radcliffe et al., 1998; Kemme et al., 1999; Boling et al., 2000). These organic acids exert their effect by decreasing stomach pH (Radcliffe et al., 1998)

High intrinsic phytase feed ingredients

Organic acids

Dietary determinants of phytase efficacy

Combined microbial phytases

Combined feed enzymes

Fig. 9.3. Determinants of dietary phytase efficacy.

Liquid feeding

Low-phytate crops

Phytase production systems

Plasma phosphate (mg/dl)

Animal Nutrition

7 6

1:1 2:1

5 4 3 Day 0

Day 10

Day 20

Day 30

Fig. 9.4. Effect of dietary calcium/phosphorus ratio on plasma inorganic phosphate concentrations of pigs fed 750 units of phytase/kg of maize–soybean meal diet. (From Lei et al., 1994.)

and lowering the rate of gastric emptying. Interestingly, addition of these acids in phytase (Fig. 9.5) or vitamin D-supplemented diets produces additive or synergistic effects on the individual action of these components on nutrient utilization (Han et al., 1998; Kemme et al., 1999; Maguire et al., 2003; Snow et al., 2004).

Augmented Effects of Microbial Phytase and Other Strategies Feed ingredients of wheat, barley, oat and their co-products contain relatively high intrinsic phytase activity (Eeckhout and De Paepe, 1994; Viveros et al., 2000; Zimmermann et al., 2002;

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Shen et al., 2005). Inclusion of these ingredients in swine diets with sufficient phytase activity can therefore supply adequate phosphorus nutrition for the growing period (Pointillart et al., 1987) or the entire growing to finishing period (Han et al., 1997). The amount of microbial phytase activity needed for swine and poultry diets can be reduced by including high intrinsic phytase ingredients such as wheat middlings (Han et al., 1998) or barley (Skoglund et al., 1998; Carlson and Damgaard Poulsen, 2003). However, synergistic effects have not been observed from the combination of various microbial phytases (Stahl et al., 2001, 2004; Augspurger et al., 2003; Gentile et al., 2003). This is intriguing, because phytases with distinctly different initiation site, substrate affinity, pH profile and proteolysis resistance may be functionally complementary. Appropriate conditions for promoting the possible synergistic functions of different phytases should be further explored. Combined supplementation of phytase with other feed enzymes such as carbohydrases or proteases has been employed to improve the overall nutrient utilization of animal feeds. The combination of xylanase, glycanase, or glycosidase with phytase showed additive effects on phytate digestibility (Zyla et al., 1999a), apparent metabolizable energy (Ravindran et al., 1999b; Wu et al., 2004) and growth (Zyla et al., 1999b; Juanpere et al., 2005). Fungal acid protease and cellulase promoted phytase-mediated dephosphorylation of myo-inositol hexakisphosphate in vitro

a

Weight gain (g/day)

a 500

b 400

Inorganic P (%) Wheat middlings (%) Microbial phytase (units/kg) Citric acid (%)

0.2 0 0 0

0 15 300 0

0 15 300 1.5

Fig. 9.5. Interaction among dietary microbial phytase, plant phytase and citric acid on daily weight gain of pigs. (From Han et al., 1998.) a vs. b: P < 0.05

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(Zyla et al., 1995). Feeding swine or poultry with multiple enzyme preparations (Omogbenigum et al., 2004) improved growth performance and digestibility of phytate and other components. Based on growth performance and serum criteria, Liu et al. (1997) observed that 250 units of phytase/kg diet with liquid feeding was as effective as 500 units/kg with dry feeding of a maize–soybean meal diet. In addition, soaking plus phytase tended to improve phosphorus and calcium digestibility in comparison with dry feeding plus phytase, thus reducing the amount of faecal phosphorus excretion. Skoglund et al. (1998) found that supplementation of microbial phytase to a barley–rapeseed diet and steeping in whey for 3 h at 40°C reduced faecal phytate content in pigs by 64%. There was a novel application of liquid feeding in which phytate was degraded by phytase from dietary supplementation, intrinsic sources of feed ingredients and production by the fermenting microbes. Lactic acid was generated from the fermentation of liquid feed, and the population of lactic acid bacteria in the gastrointestinal environment of the pig was increased (Canibe and Jensen, 2003; Carlson and Damgaard Poulsen, 2003). Low-phytate varieties of maize (Ertl et al., 1998) and barley (Hatzack et al., 2000) have comparable levels of total phosphorus to the parental lines, but greater bioavailability of phosphorus to swine and poultry (Ertl et al., 1998; Overturf et al., 2003; Thacker et al., 2004). The potential of these low-phytate varieties for improving feed phosphorus utilization can be further augmented by phytase supplementation (Yan et al., 2000; Applegate et al., 2003a; Baxter et al., 2003). Meanwhile, transgenic soybean and canola lines that overexpress the phyA gene of A. niger have been developed, and lead to improved utilization of phosphorus and calcium by poultry (Denbow et al., 1998; Zhang et al., 2000). Onyango et al. (2004) suggested that differences in glycosylation of E. coli phytase among yeast production systems might affect the enzyme efficacy. In contrast, Zhang et al. (2000) found no difference in efficacy of A. niger PhyA phytase derived from a microbial or plant (canola) expression system.

Constraints of Phytase Application Stability and cost are the two major constraints for the application of currently available phy-

tases. An ‘ideal’ phytase should be stable for feed pelleting and storage. The high temperatures used in the pelleting process denature phytase and thus reduce the enzyme activity in the final products (Wyss et al., 1998; Igbasan et al., 2000). Phytase coating has been developed to counteract this destructive effect of thermal processing, but does not ensure a complete release of the enzyme from the granules during its transit through the gastrointestinal tract of the animal. Spraying of liquid phytase to feed post pelleting has been used to bypass phytase denaturation by pelleting, but equipment costs are substantial. It is also challenging to ensure an even distribution of phytase by spraying (Johnston and Southern, 2000). The phytase product should be stored under dark, dry and cool conditions. Refrigeration of the enzyme will extend its shelf life, and powder preparations are usually more stable than liquid preparations. If possible, phytase should be stored apart from mineral and vitamin premixes. The economic returns of phytase application are determined by the price of the phytase product and the equipment needed for phytase supplementation, in relation to the amounts of non-phytate phosphorus, calcium and other dietary components that can be spared by the use of phytase. Thus, application of phytase is economically attractive in places where strict regulations impose fines to the excess of animal manure phosphorus. Phytase is generally considered safe for handlers or users, and only minor allergic reactions have been described among workers of a technical centre for large-scale confection of phytase in powdered form (Baur et al., 2002).

Developing Thermostable Phytases A thermostable phytase has been isolated from A. fumigatus. Compared with phytases isolated from A. terreus, Myceliophthora thermophila or A. niger (Pasamontes et al., 1997; Wyss et al., 1998), this phytase is more thermotolerant. This feature of the enzyme can be modulated by specificity of buffers used in the heat treatments (Rodriguez et al., 2000a) and appears to be related to its ability to refold after heat denaturation (Wyss et al., 1998). However, there is no major difference in its three-dimensional structure from that of A. niger PhyA (Xiang et al., 2004). Developing thermostable enzymes capable of withstanding feed pelleting has been intensively

Animal Nutrition

studied. State of the art for this purpose includes directed evolution, in which repetitive rounds of in vitro diversification are conducted and tested using high throughput screening methodologies. Garrett et al. (2004) optimized the performance of E. coli AppA phytase by gene site saturation mutagenesis. They generated a library of single site mutations of the individual amino acid residues of the enzyme. After identification of all single point mutations that enhanced the thermal stability of the enzyme, a new screen was designed to discriminate between the most stable of the combinatorial products using a sequential, recursive addition protocol. As a result, a final construct termed Phy9X was developed with significantly higher thermostability and gastric resistance than the parent AppA enzyme. Rational design is another powerful tool for the development of thermostable phytases. Based on the role of glycosylation on A. niger PhyA stability (Han et al., 1999), Rodriguez et al. (2000b) designed several mutations to add potential glycosylation sites in E. coli AppA phytase expressed in the methylotrophic yeast Pichia pastoris. Two of the mutations showed elevated glycosylation. However, the increased glycosylation did not affect thermostability of the mutated enzymes. Another mutant did not show a higher degree of glycosylation, but did show a higher thermostability and improved kinetic properties compared with the wild-type enzyme. These changes were attributed to a higher number of hydrophobic interactions and disappearance of a disulphide bond between a G helix and GH loop present in the α-domain of the protein that would improve its flexibility and catalytic properties. The feeding efficacy of this mutant phytase to release phosphorus from phytate in a maize–soybean diet was tested in a pig trial (Gentile et al., 2003). However, Wyss et al. (1999) did not observe any effect of glycosylation on thermostability of phytase. Based on primary protein sequence analysis of fungal phytases, Lehmann et al. (2000a) chose the most conserved amino acid residues and constructed a novel phytase termed consensus phytase-1. The synthetic enzyme featured a higher optimum temperature (16–29°C increase) and unfolding temperature (15–22°C) when compared to the parent phytases, but no change in their catalytic properties. Stabilization of the consensus phytase resulted from a combination of several amino acid residues that were located mainly in regions with no defined secondary

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structure on the surface of the molecule (Lehmann et al., 2000b, 2002). The consensus phytase was effective in releasing phosphorus from phytate in diets for swine or poultry (Gentile et al., 2003; Paditz et al., 2004; EsteveGarcia et al., 2005). In addition, Jermutus et al. (2001) expanded the development of structurebased chimeric enzymes to the replacement of an entire α-helix on the surface of A. terreus phytase.

Enhancing Proteolysis Resistance of Phytases Resistance to proteolysis is of importance for phytase to function in the digestive tracts of animals. Given that phytate hydrolysis mainly takes place in pig stomach or the crop, gizzard and proventriculus of poultry, resistance to pepsin, the main protease in situ, is desirable for efficient phytases. Rodriguez et al. (1999a) found a greater pepsin resistance of the recombinant AppA phytase from E. coli expressed in P. pastoris than that of the recombinant A. niger PhyA phytase. In fact, the activity of E. coli phytase was elevated after the pepsin digestion. Resistance of E. coli AppA to pepsin digestion has been further confirmed by Golovan et al. (2000). Simon and Igbasan (2002) found a higher susceptibility to proteolytic cleavage of several fungal phytases when compared to a bacterial or consensus phytase. On the other hand, A. niger PhyA phytase exhibited a lower susceptibility to proteolysis when compared to wheat or yeast phytases (Phillippy, 1999; Matsui et al., 2000). However, A. fumigatus phytase was highly susceptible to trypsin (Rodriguez et al., 2000a). Kim et al. (2003) described pepsin and trypsin resistances of a novel phytase isolated from Citrobacter braakii. Proteolysis resistance of phytase may be enhanced by site-directed mutagenesis that modifies cleavage sites of exposed loops susceptible to protease action (Wyss et al., 1999). Modifications of E. coli AppA phytase by gene site saturation mutagenesis caused a 3.5-fold enhancement in gastric stability of the enzyme (Garrett et al., 2004). Gastrointestinal carriers may also be used to help phytase in reducing stomach and small intestine proteolytic degradation. Haraldsson et al. (2005) reported the use of high phytase-producing strains of Saccharomyces cerevisiae as the phytase carriers. The limitation of this approach is the pH

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dependence of phytase expression by the yeast, and the narrow pH profile of the phytase activity. This could be partially overcome by the use of genetically modified yeast species that produce recombinant phytases active in the acidic milieu of the stomach.

Improving the pH Profile and Catalytic Properties of Phytases Isolation and cloning of phytases with higher catalytic efficiency than the currently commercialized fungal phytases have been a focus of phytase research. AppA phytase isolated from E. coli and expressed in P. pastoris production system was as efficient as, or superior to, A. niger PhyA phytase (Stahl et al., 2000). A novel E. coli phytase gene (appA2) was isolated from pig colon with a 95% sequence homology to appA, but the expressed phytase in P. pastoris had different biochemical properties (Rodriguez et al., 1999b). As mentioned above, other groups showed the superior effectiveness of this AppA2 phytase to that of the commercialized phytases from Peniophora lycii or A. niger in diets for swine or poultry (Augspurger et al., 2003; Applegate et al., 2003a). Most of the mutation studies for improving the pH profile of phytase have been done using A. niger or A. fumigatus phytase. As summarized by Tomschy et al. (2002), three major approaches can be taken for the purpose: 1. modification of ionizable groups directly involved in substrate specificity or catalysis; 2. replacement of amino acid residues in direct contact with residues located in the active or substrate specificity site by means of hydrogen bonds or salt bridges; 3. alteration of distant charge interactions by modification of the surface charge of the enzyme. Tomschy et al. (2002) improved the activity at low pH of A. fumigatus phytase and a consensus phytase. Their approach included decreasing surface charge through glycinamidylation and replacement of active site residues using site-directed mutagenesis based on sequence alignments and experimentally determined or homologically modelled three-dimensional structures of A. niger

PhyA phytase, P. lycii phytase or A. niger pH 2.5 acid phosphatase. Using molecular modelling of the substrate specificity site of A. niger phytase and sequence comparisons, Mullaney et al. (2002) successfully developed a recombinant phytase with the substitution of lysine by glutamic acid at position 300, which enhanced the specific activity of the enzyme at pH levels between 4 and 5. The authors suggested that the residue substitution would lower the local electrostatic field attraction for myo-inositol hexakisphosphate at both pH ranges and thus enhance the enzyme kinetics. Recently, our laboratory (Kim et al., 2006) produced a shift of optimal pH of A. niger PhyA phytase from 5.5 to 3.5–4.0 by altering the charges of amino acid residues in the substrate binding site. The variant showed a significant enhancement of function in the stomach of pigs. By substituting the glutamic acid residue located at position 27 by leucine, as in A. terreus phytase, Tomschy et al. (2000a) improved the specific activity of A. fumigatus phytase without changing its substrate specificity. The improvement appeared to be due to a weakening or loss of a hydrogen bond between glutamic acid at position 27 and the 6-phosphate group of myoinositol hexakisphosphate, suggesting that product release would be the rate-limiting step of the A. fumigatus phytase reaction. In a different set of experiments, Tomschy et al. (2000b) improved the catalytic properties of A. niger T213 phytase by site-directed mutagenesis of R297Q. A. niger T213 phytase has 12 divergent amino acids from those of A. niger NRRL 3135 phytase. Of these divergent amino acids, three are located in the vicinity of the active site and were substituted by their divergent counterparts of A. niger NRRL 3135 phytase. The R297Q substitution improved specific activity and pH profile of A. niger T213 phytase to levels comparable to A. niger NRRL 3135 phytase. As the wild-type A. niger T213 phytase had a lower specific activity for myo-inositol hexakisphosphate, but not for other smaller and/or less negatively charged phosphate substrates, they suggested that product (myo-inositol pentakisphosphate) release was the rate-limiting step in the enzyme reaction due to interaction of the guanidino group of arginine in position 297 of A. niger T213 phytase with one of the phosphate groups of myo-inositol pentakisphosphate.

Animal Nutrition

To improve the catalytic properties of the thermostable consensus-1 phytase, Lehmann et al. (2000b) replaced all the divergent amino acid residues present in the active site of the consensus phytase by those of A. niger NRRL 3135 phytase. The new phytase, termed consensus-7 phytase, featured a major shift in the catalytic properties that were similar to those of A. niger NRRL 3135 phytase, thus demonstrating the feasibility of rational transfer of favourable catalytic properties. However, the active site residues transfer caused a decrease in the unfolding temperature of consensus-7 phytase compared with consensus-1 phytase.

The Search for Efficient Phytase Production Systems Selection of the phytase expression system is crucial to ensuring an efficient and affordable enzyme production. The expression system of choice may vary with the origin and properties of the phytase to be expressed. Because transgenic plants appear to be a cost-effective fermenter for biological materials, successful attempts to overexpress microbial phytases have been made in soybean, wheat, rice, lucerne and canola (Li et al., 1997; Brinch-Pedersen et al., 2000; Ponstein et al., 2002; Ullah et al., 2002; Hong et al., 2004). The effectiveness of phytases produced in these plants have been determined (Pen et al., 1993; Denbow et al., 1998; Zhang et al., 2000; Hong et al., 2004), but practical problems include public concern over genetically modified organisms and the relatively low thermostability of the expressed phytases in plants (Igbasan et al., 2000; Lucca et al., 2001). Fungal systems from the Aspergillus genera (A. niger, A. oryzae and A. awamori) are employed for phytase production (Mitchell et al., 1997; Martin et al., 2003). Several strategies have been used to avoid proteolysis associated with Aspergillus expression systems, including supplementation of a complex inducing medium with increasing concentrations of yeast autolysate or additional protein sources like malt extract or casamino acids. Methylotrophic yeast such as P. pastoris or Hansenula polymorpha combine several appealing features such as efficient post-translational modifi-

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cation, capability of secreting extremely high levels of recombinant protein, ease of manipulation and rapid growth. The enzyme production can be greatly enhanced by optimizing fermentation conditions (Mayer et al., 1999; Chen et al., 2004). Rodriguez et al. (1999a,b, 2000a,b) reported efficient heterologous expression of A. niger, E. coli and A. fumigatus phytase in P. pastoris using inducible expression driven by the potent alcohol oxidase promoter (AOX1). Lee et al. (2005) expressed the AppA2 phytase using both inducible and constitutive expression systems, and obtained lower yields in S. cerevisiae and Schizosaccharomyces pombe than in the P. pastoris expression systems. Never the less, the expression systems or hosts had no effect on biochemical properties of the recombinant phytases. These similarities offer flexibility for choosing phytase fermentation systems. Several bacterial expression systems have been studied for phytase expression. To avoid extensive manipulation and purification steps due to periplasmic expression of recombinant proteins by E. coli, Miksch et al. (2002) tested extracellular expression of E. coli phytase using a secretion system based on the controlled expression of the kil gene. They analysed major factors such as promoter type, host strain and selection pressure that could affect the level of heterologous expression, and developed an effective fedbatch fermentation strategy. Kerovuo et al. (2000) studied a novel Bacillus expression system and demonstrated its potential application for efficient extracellular phytase production, whereas Stahl et al. (2003) attempted extracellular expression of E. coli AppA phytase in Streptomyces lividans and studied changes in its biochemical properties when compared to a glycosylated AppA produced by P. pastoris. Bacillus subtilis phytase has been expressed in Lactobacillus plantarum 755 (Kerovuo and Tynkkynen, 2000). Despite the low expression and secretion levels, the authors suggested that culture conditions could be optimized for a higher yield. In addition, the bacterial strain could be used as an inoculum for fermented plant material or as phytase carrier in combination with the probiotic effects inherent to lactic acid bacteria. In general, bacterial expression systems have a disadvantage compared to plant, fungal or yeast expression systems for their inability to

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glycosylate the recombinant protein expressed. Glycosylation has been shown to be important for fungal phytase expression (Han and Lei, 1999). Phillippy and Mullaney (1997) found that when the A. niger phytase gene was expressed intracellularly in E. coli, the expressed protein was inactive. Yin et al. (2005) recently described a novel phytase expression system based on silkworm larvae infected with baculovirus transfected with the baculovirus transfer vector pVL

1393 under the control of the polyhedron promoter and harbouring the E. coli appA phytase gene. Using this system, the authors obtained a phytase yield of 7710 units/ml hemolymph.

Future Perspectives Figure 9.6 depicts strategies and paths for developing ‘ideal’ phytases that are catalytic-efficient,

Biotechnology

Resistance to proteolysis

High intrinsic resistance to proteolysis

Catalysis and pH profile

Site-directed mutagenesis Phytase carriers

New phytase genes with higher intrinsic activity Chemical modification Site-directed mutagenesis Directed evolution Exchange of active site

Thermostability

Naturally thermostable enzymes

Directed evolution

Consensus concept for thermostability

Rational design

Structure-based chimeric enzymes

Development of more efficient production systems

Transgenic plants and animals

Fungal systems Aspergillus spp.

Yeast systems Saccharomyces cerevisiae Pichia pastoris Hansenula polymorpha Schyzosaccharomyces pombe

Bacterial systems Escherichia coli Bacillus spp. Streptomyces lividans Lactobacillus plantarum

Novel expression systems Baculovirus-infected silkworm larvae Fig. 9.6. Biotechnology for the development of catalytic-efficient, heat-stable, proteolysis-resistant and economical phytases.

Animal Nutrition

protease-resistant, heat-stable and cost-effective for different species at various physiological stages. The success of this attempt will overcome the current constraints of phytase application. A wide application of phytase in animal diets, together with other nutritional and environmental measures, will help alleviate or eliminate the manure phosphorus pollution problem worldwide. Transgenic plants overexpressing phytase in leaves and seeds have recently been developed and may offer another economic source of phytase for animal or human nutrition. Meanwhile, lowphytate grains may provide an appreciable level of available phosphorus to animals. Transgenic pigs (Golovan et al., 2001) with overexpressed phytase in their saliva require lower levels of

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inorganic phosphorus supplements in plant-based diets and excrete ~70% less of faecal phosphorus than the controls. Hopefully, these novel technologies can be combined to effectively improve the utilization of dietary phytate by animals and minimize environmental phosphorus pollution from their manure.

Acknowledgements The phytase research in Xin Gen Lei’s laboratory was funded in part by the Cornell Biotechnology Program. Dr Porres works under a research contract from Junta de Andalucia, Spain, and Project AGL2002-02905 ALI.

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10

Environmental Implications of Inositol Phosphates in Animal Manures April B. Leytem1 and Rory O. Maguire2 1

United States Department of Agriculture–Agricultural Research Service, Northwest Irrigation and Soils Research Laboratory, 3793 N. 3600 E., Kimberly, ID 83341, USA; 2Crop and Soil Environmental Sciences, Virginia Tech, Box 0404, Blacksburg, VA 24061, USA

Animal production in the USA is valued at more than $100 billion and has consolidated significantly during the last 20 years, with a larger number of animals being produced on an increasingly smaller land base (Kellogg et al., 2000). Manure generated from animal production is currently estimated to exceed 335 million t of dry matter per year in the USA, while global manure production is estimated at ~13 billion t of dry matter per year (Mullins et al., 2005). Manures contain significant amounts of phosphorus, with values between 6.7 and 29.1 g P/kg on a dry weight basis reported for several species of animals (Barnett, 1994). This phosphorus includes inorganic and organic forms, with the latter constituting between 10% and 80% of the total (Peperzak et al., 1959; Gerritse and Zugec, 1977). Inositol phosphates are one of the primary organic phosphorus species found in manures, with myo-inositol hexakisphosphate typically being the most abundant (Peperzak et al., 1959; Barnett, 1994; Turner and Leytem, 2004). The environmental fate of phosphorus in animal manures is determined in part by the chemical composition of the phosphorus, yet few studies have fully characterized manure phosphorus and determined the effect of the various phosphorus compounds on phosphorus behaviour in soil. The various forms of organic phosphorus differ in the extent of their sorption when applied to soils, with myo-inositol hexakisphosphate being strongly bound while other organic phosphorus 150

compounds such as nucleotides, DNA and glucose phosphates are more mobile (Celi and Barberis, 2005). Phosphorus applied to soil as manure may also behave differently from mineral phosphate fertilizer, due to other chemical characteristics of the manure. Organic matter in manure can complex iron and aluminium via organic ligands, which decreases the precipitation of inositol phosphates with these metals. It also competes for sorption sites in soil, increasing the concentration of phosphate in solution (Iyamuremye et al., 1996). Inositol phosphates in manure can also disperse soil colloids and therefore increase the potential for particulate phosphorus transport in runoff (see Celi and Barberis, Chapter 13, this volume). Based on this evidence, more detailed information on the forms of phosphorus in manures, as well as those manure characteristics that influence phosphorus sorption, may shed light on the potential for off-site losses of phosphorus from land application of manure. This chapter addresses environmental issues concerning phosphorus and inositol phosphates in animal production. We summarize studies on the phosphorus composition of manures, including those using traditional extraction procedures and the more recent application of nuclear magnetic resonance (NMR) spectroscopy. Finally, we review how dietary modification and storage alters the phosphorus composition of manures, and explore the impact of such alterations on

©CAB International 2007. Inositol Phosphates: Linking Agriculture and the Environment (eds B.L. Turner, A.E. Richardson and E.J. Mullaney)

Environmental Implications in Animal Manures

phosphorus solubility in soils and the potential for phosphorus transfer in runoff.

Why is Manure Phosphorus an Environmental Concern? Consolidation of animal production can generate regional and farm-scale nutrient surpluses where nutrient imports in feed and mineral fertilizer exceed nutrient exports in crops and animal products (Sharpley et al., 1994; Sims et al., 1998). These nutrient surpluses can in turn increase the risk of nutrient loss to the environment and pollution of water bodies (Sharpley, 1996; Sims et al., 1998, 2000). Nutrients in manures can be recycled by application to cropland, which reduces the need for commercial fertilizers. Unfortunately, large amounts of manure produced in localized areas, coupled with the high cost of effective nutrient utilization strategies in an unbalanced system, favour manure disposal via land application in excess of crop nutrient needs, rather than utilizing manure in areas with nutrient deficiencies (Sharpley et al., 1998). Phosphorus is a particular concern, because it can accumulate in soil to concentrations greater than those needed for optimum crop production. This is due in part to unfavourable nitrogen/ phosphorus ratios in manures relative to the uptake of these nutrients by most crops, which results in overapplication of phosphorus when manures are applied to meet the nitrogen requirement of the crop (Mikkelsen, 2000). As a result, long-term manure application to agricultural land leads to soil phosphorus accumulation and greater potential for phosphorus transfer in runoff to water bodies. This can contribute to eutrophication in freshwater ecosystems, and numerous examples of water quality impairment associated with phosphorus pollution from animal operations now exist (Burkholder and Glasgow, 1997; US Geological Survey, 1999; Boesch et al., 2001). There is therefore an urgent need to understand and reduce the impact of animal manures on the pollution of water bodies. This demands a mechanistic understanding of the behaviour of manure phosphorus in soils and its potential for phosphorus transfer in runoff. Important aspects include the manure character-

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istics that determine phosphorus behaviour following land application and the potential changes induced by dietary modification.

Phosphorus Composition of Animal Manures Investigation of the dynamics of manure phosphorus following application to soils requires information on the phosphorus composition of the manure. One of the earliest studies of manure characterization was performed by Funatsu (1908), who used sequential extraction techniques to fractionate the phosphorus in guano. The procedure involved dilute acid to extract inorganic phosphate, inositol phosphates and other organic forms, followed by ether and alcohol to extract phospholipids, with the residue (unextracted fraction) being labelled as nucleic acid. Variations of this procedure were subsequently used by others to characterize manures from pigs fed a variety of feed rations (Rather, 1918), poultry and mixed farmyard manure (Ghani, 1941), sheep manure (McAuliffe and Peech, 1949) and fresh manure from horses, cattle, sheep, pigs and hens (Kaila, 1948). Organic phosphorus in these studies ranged between 18% and 50% of the total phosphorus, with the acidsoluble organic phosphorus (which typically included inositol phosphates) constituting between 0% and 86% of the total organic fraction. Peperzak et al. (1959) used a similar sequential extraction procedure to determine the phosphorus composition of a variety of manures. Total phosphorus concentrations ranged between 4 and 30 g P/kg dry weight, with the inorganic fraction constituting 53–95% of total phosphorus (Table 10.1). In this procedure, myo-inositol hexakisphosphate was isolated from the acid extract and was found to represent between 1% and 22% of total phosphorus, with other acid-soluble organic phosphorus forms constituting between 3% and 44%. The alcohol-soluble fractions were small (0.4–1.3%) while residual phosphorus values ranged between 2% and 27% of total phosphorus. When manures of different ages were examined from a stockyard, the general trend was a decrease in organic phosphorus from 49% to 32% of total phosphorus over 20 years, with a

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A.B. Leytem and R.O. Maguire

Table 10.1. Concentrations of phosphorus compounds in sequential extracts of animal manures. (From Peperzak et al., 1959.)

Phosphate Animal

Total P (g P/kg)

Chick Hen Sheep Sow Horse Steer Bull Cow Calf

13–23 7–30 12 11 4–7 8–12 9 4–7 5

myo-Inositol hexakisphosphate

Other acidsoluble P

Alcoholsoluble P

Residual P

0.6–1.0 0.1–0.6 0.4 0.5 0.8 1.0 0.7–1.0 1.3 0.4–1.3

2–27 5–12 16 3 2–20 13–19 14 3–14 17

% of total P 53–56 54–81 63 83 73–95 60–64 76 67–87 62

NDa 12–22 2 0.6 1–2 7–10 0.5 1–5 3

17–44 3–11 19 13 14 12–13 8 7–25 17

a

ND = not detected.

concomitant decrease in myo-inositol hexakisphosphate from 3.9% to 1.5% of total phosphorus. Barnett (1994) published the most recent comprehensive study on organic phosphorus compounds in animal manures using conventional sequential fractionation techniques. Organic phosphorus in a variety of manures was fractionated into phospholipids, nucleic acids, acid-soluble organic phosphorus, inorganic phosphate and residual phosphorus. Inorganic phosphate constituted the greatest proportion of the total phosphorus, followed in descending order of magnitude by residual phosphorus, acid-soluble organic phosphorus and small amounts of phospholipids. In this study the myo-inositol hexakisphosphate content was not directly measured, but the acid-soluble organic phosphorus fraction, which typically includes the inositol phosphates, ranged between 7.8% and 53.4% of the total phosphorus. Interest in the environmental fate of manure phosphorus prompted recent studies to adopt the Hedley fractionation (Dou et al., 2000; Sharpley and Moyer, 2000; Weinhold and Miller, 2004). This procedure was originally developed to assess phosphorus solubility in soil (Hedley et al., 1982) and involves sequential extraction with water, sodium bicarbonate, sodium hydroxide and hydrochloric acid. Phosphorus extracted in water and bicarbonate is considered readily soluble, while that extracted in sodium hydroxide (assumed to be associated with amorphous iron/aluminium and organic matter) and hydrochloric acid

(assumed to be calcium phosphates) is considered poorly soluble. However, several problems compromise the suitability of the Hedley fractionation for manures. In particular, phosphorus chemistry differs markedly between soils and manures, being controlled commonly by iron and aluminium oxides and calcium carbonate in soils (Hedley et al., 1982), and by association with calcium and magnesium in manures (Cooperband and Ward Good, 2002). Turner and Leytem (2004) used solution 31P NMR spectroscopy to unequivocally identify phosphorus compounds in the various fractions of the Hedley extraction scheme as applied to poultry, swine and cattle manures. Two main groups of phosphorus compounds were determined with this procedure: a readily soluble fraction extracted with water and sodium bicarbonate and a stable fraction extracted with sodium hydroxide and hydrochloric acid. Organic phosphorus in the readily soluble fraction included DNA, phospholipids and simple phosphate monoesters. Organic phosphorus in the stable fraction consisted mainly of myo-inositol hexakisphosphate. Since there was considerable overlap between the extracts, the authors recommended a simpler procedure consisting of extraction with sodium bicarbonate to remove the readily soluble fraction (which would be most susceptible to transport in runoff), followed by extraction with a solution containing sodium hydroxide and ethylenediamine tetraacetate (EDTA) to recover the more stable

Environmental Implications in Animal Manures

fraction. This method gave near-quantitative recovery of phosphorus from swine and poultry manure (Turner, 2004; Turner and Leytem, 2004). Solution 31P NMR spectroscopy has been used to quantify the phosphorus composition of a wide variety of manures (Leinweber et al., 1997; Leytem et al., 2004; Maguire et al., 2004; Turner, 2004; Turner and Leytem, 2004; McGrath et al., 2005). These studies indicate that manure phosphorus is predominately inorganic phosphate, followed in descending order by phosphate monoesters, phosphate diesters (nucleic acids and phospholipid), pyrophosphates and, in some cases, phosphonates. Concentrations of myo-inositol hexakisphosphate ranged from non-detectable to 80% of the total phosphorus in manures from a variety of ruminant (cattle and sheep) and monogastric animals (poultry, swine; Table 10.2). Solidstate 31P NMR spectroscopy has also been applied to manures (e.g. Hunger et al., 2004), but cannot accurately assess the organic phosphorus fraction. As demonstrated by both sequential fractionation and solution 31P NMR spectroscopy, the myo-inositol hexakisphosphate content of manures can vary widely, both among and within species (Table 10.2). There are physiological differences between ruminant and monogastric animals that can account for these differences. The diets of monogastric animals often include large amounts of cereal grains, in which much of the phosphorus occurs as salts of myo-inositol hexakisphosphate (phytate); for example, approximately two-thirds of the phosphorus in maize and soybeans is in this form (see Raboy, Chapter 8, this volume). As monogastric animals do not possess ample gut phytase (McCuaig et al., 1972), manures from poultry and pigs can contain large amounts of undigested phytate (although see Leytem et al., 2004). In contrast, ruminant animals have the capacity to hydrolyse inositol phosphates in their diet, and manures from animals fed grass or lucerne-based diets contain little phytate. However, there is evidence that for ruminants fed a grain-based diet, metal complexation can prevent extensive hydrolysis of myo-inositol hexakisphosphate and allow it to pass through the animal intact (see Dao, Chapter 11, this volume). Dietary effects are also evident within a given species. For example, manure from laying hens fed maize with varying levels of non-phytate

153

phosphorus, with and without phytase additions, can contain a wide range of myo-inositol hexakisphosphate concentrations (35–80% of total phosphorus, whereas manure from broilers fed a diet consisting mainly of barley contains closer to 10% of total phosphorus in this form (Table 10.2). This indicates the importance of determining dietary impacts on the composition of manure phosphorus excreted from the animal to assess the potential behaviour of manure phosphorus once applied on land. Since it has been demonstrated that inositol phosphates can sorb strongly to soils (see Celi and Barberis, Chapter 13, this volume), changes in the concentration of myo-inositol hexakisphosphate in manure could be of concern from an environmental standpoint (discussed later).

Impact of Dietary Manipulation on myo -Inositol Hexakisphosphate in Manure As monogastric animals cannot fully utilize phytate in cereal grains, mineral phosphate supplements are commonly added to their diets to prevent phosphorus deficiency. As described above, this increases phosphorus concentrations in manure and can lead to phosphorus accumulation in soils when manure phosphorus is applied in excess of crop phosphorus removal (Sims et al., 2000). To address concerns regarding surplus phosphorus in manure, strategies involving dietary manipulation are being widely adopted to reduce manure phosphorus concentrations (see Lei and Porres, Chapter 9, this volume). By reducing phosphorus excretion, manures with nitrogen/ phosphorus ratios more closely matching the nutrient needs of crops can be generated, thereby reducing overapplication of phosphorus and build-up of soil phosphorus. For monogastric animals that have a limited ability to digest phytate, dietary strategies include the isolation of mutant grains that store most of the total phosphorus in the grain as inorganic phosphate and less as phytate (Raboy et al., 2000; Dorsch et al., 2003, see Raboy, Chapter 8, this volume), thereby enhancing phosphorus uptake by the animal and reducing the excreted phosphorus (Spencer et al., 2000; Veum et al., 2002; Jang et al., 2003; Klunzinger et al., 2005). Supplementation of animal feeds with microbial phytase is

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A.B. Leytem and R.O. Maguire

Table 10.2. Concentrations of phosphorus compounds in extracts of manures from a selection of animals determined by solution 31P NMR spectroscopy. (From Leytem et al., 2004, 2005, 2006 and unpublished data; Maguire et al., 2004.) Total phosphorusa Manure Swine manure, fresh (barley feed) Swine lagoon liquid Broiler manure (barley feed) Broiler manure (standard maize diet) Broiler manure (maize, low NPPc) Broiler manure (maize, low NPP + phytase) Broiler litter (maize, high NPP) Broiler litter (maize, high NPP + phytase) Turkey litter (maize, high NPP) Turkey litter (maize, high NPP + phytase) Dairy lagoon liquid Dairy compost Beef manure (maize-fed) Beef manure (pasture-fed) Sheep (barley-fed)

Phosphateb

Phosphate monoestersb

Pyrophosphateb

myo-Inositol hexakisphosphateb

13.46 (97)

13.02 (94)

0.67 (5)

0.13 (1)

Tr

30.00 (99)

29.15 (97)

0.75 (3)

0.09 (<1)

ND

6.36 (99)

4.46 (70)

1.92 (30)

ND

0.74 (12)

15.61 (96)

7.21 (46)

8.19 (53)

0.21 (1)

7.61 (49)

9.49 (99)

1.22 (28)

8.17 (86)

0.10 (1)

7.62 (80)

9.61 (98)

5.33 (56)

4.04 (42)

0.13 (1)

3.39 (35)

13.90 (98)

5.71 (41)

8.38 (60)

0.06 (<1)

7.83 (56)

10.40 (96)

5.05 (49)

5.74 (55)

ND

4.88 (47)

15.40 (87)

10.90 (71)

6.74 (44)

0.14 (1)

5.09 (33)

12.80 (94)

8.56 (67)

4.82 (38)

0.14 (1)

3.45 (26)

8.80 (93) 2.50 (98) 4.20 (99)

7.93 (90) 2.28 (91) 2.51 (60)

0.82 (9) 0.22 (9) 1.60 (38)

0.06 (<1) 0.004 (<1) 0.09 (2)

0.37 (4.2) 0.03 (1) 0.34 (8)

4.10 (83)

2.65 (65)

1.0 (25)

0.25 (6)

ND

8.45 (91)

5.52 (65)

1.68 (20)

0.41 (5)

0.47 (6)

g P/kg dry wt

a

Values are total phosphorus extracted by sodium hydroxide and ethylenediaminetetraacetate (EDTA), and values in parentheses are the proportion (%) of the total manure phosphorus determined by microwave digestion. b Values in parentheses are the proportion (%) of the extracted phosphorus. c NPP = non-phytate phosphorus. Tr = trace; ND = not detected.

also used to increase phytate hydrolysis in the gut, thereby enhancing phosphorus utilization by the animal (Cromwell et al., 1993; Coelho and Kornegay, 1996; see Lei and Porres, Chapter 9, this volume). The combination of low-phytate

grains with phytase additions is also utilized to further reduce phosphorus excretion. In addition to reducing the concentrations of phosphorus in manure, dietary modification is expected to influence manure phosphorus com-

Environmental Implications in Animal Manures

position, which may have implications for the environmental fate of manure phosphorus (Turner et al., 2002). Potentially the greatest impact of diet modification in monogastric animals on phosphorus forms in manure is likely to be changes in the amount of phytate excreted, with a corresponding increase in the proportion of the manure phosphorus that occurs as waterextractable phosphate. Thus, as diet modification reduces the proportion of the manure phosphorus occurring as myo-inositol hexakisphosphate, the proportion of water-extractable phosphate in the manure increases as a fraction of total phosphorus, even though the total phosphorus concentration may be reduced. This is particularly evident for poultry manures (Fig. 10.1) and may be important when manures are applied to land on the basis of phosphorus content, as is now common in several states in the USA. Feeding low-phytate grains

Manure water-extractable P (% total P)

Mutant grains that contain substantially less phytate than the wild-type equivalent that has traditionally been fed to animals (Raboy et al., 2000; Dorsch et al., 2003; see Raboy, Chapter 8, this volume) have recently been developed. At present there are low-phytate varieties of maize, barley and soybean meal that can be used in feed formulations. Low agronomic yields of these mutant grains have prevented wide adoption, but future improvements are likely, and these grains

100 Toor et al. (2005) Maguire et al. (2004) Leytem et al. (2006a)

80 60 40 20 0 0

20

40

60

80

100

Manure phytate (% total P)

Fig. 10.1. The effect of phytate concentration on water-extractable phosphorus in manures from modified poultry diets. (From Maguire et al., 2004; Toor et al., 2005; Leytem et al., 2006.)

155

will be useful for developing strategies to reduce phosphorus excretion by monogastric animals. Large reductions in total phosphorus excretion can be achieved using these grains (Spencer et al., 2000; Li et al., 2001; Veum et al., 2002; Jang et al., 2003; see Lei and Porres, Chapter 9, this volume), although only a few studies have determined their impact on phosphorus composition in manure. Toor et al. (2005) reported a decrease of only 10% in excreted total phosphorus from broilers fed diets containing normal maize vs. low-phytate maize, although there was a 47% reduction in the amount of myo-inositol hexakisphosphate excreted by the birds. Baxter et al. (2003) saw the same trend for swine fed low-phytate maize; total phosphorus excretion was only slightly reduced, but myo-inositol hexakisphosphate excretion was reduced by almost 50%. When low-phytate barleys were included in broiler diets, manure total phosphorus concentrations were reduced by 14–24% (Leytem et al., 2006b; Table 10.3). However, myo-inositol hexakisphosphate concentrations in manures from all dietary treatments constituted only 3–12% of the total phosphorus in the manure, even when as much as 91% of total phosphorus in the feed was phytate. This same trend was also reported for swine in a similar study; total phosphorus excretion was reduced by ~33% when animals were fed lowphytate diets, yet myo-inositol hexakisphosphate was excreted only in trace amounts (Leytem et al., 2004; Table 10.3). This indicates that even though monogastric animals do not possess sufficient phytase to hydrolyse phytate in the part of the digestive tract where phosphorus sorption takes place, the phytate is not necessarily excreted by the animal. A possible explanation is that barley diets contain high intrinsic phytase activity (see Lei and Porres, Chapter 9, this volume), which might lead to phytate hydrolysis in the animal. However, in a study of swine manure from animals fed diets containing wild-type and low-phytate maize, which contains little intrinsic phytase, most of the excreted phosphorus (~80% of total phosphorus) was inorganic phosphate and there was little difference in the manure fractions across dietary treatments (Weinhold and Miller, 2004). A more likely explanation, therefore, is that phytate is hydrolysed in the hindgut by intestinal microflora, even though the animals derive little nutritional benefit from this process in the lower intestine.

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Table 10.3. Phosphorus concentrations in poultry and swine manure fed either a wild-type barley (Copeland and CDC Bold) or mutant barley with reduced amounts of grain phytic acid content (M 422, M 635, M 955). Phosphorus concentrations were determined by extraction in sodium hydroxide and ethylenediaminetetraacetate (EDTA) and solution 31P NMR spectroscopy. Means in the same column (for each animal type) followed by the same letter do not differ significantly (P > 0.05). (From Leytem et al., 2004; Leytem, A.B., Thacher, P.A. and Turner, B.L., 2006, unpublished data.) NaOH–EDTA extractable P (g P/kg dry wt)

Grain type

Feed phytate (% total phosphorus)

Total Pa

Phosphateb

Phosphate monoestersb,c

myo-Inositol hexakisphosphateb

Poultry (broiler chicks) Copeland M 422 M 635 M 955

91 40 37 <1

6.36 4.48 4.93 5.15

CDC Bold M 422 M 635 M 955

55 50 26 3

13.46 8.55 8.05 8.36

(99)a 4.46 (70)a (93)c 3.42 (69)c (92)bc 3.20 (72)c (92)b 3.88 (75)b Swine (barrows)

1.92 1.53 1.29 1.24

(30)a (31)b (29)b (24)b

0.74 0.34 0.34 0.14

(97)a (95)b (91)b (95)b

0.67 1.08 1.08 0.91

(5)a (12)a (12)a (11)a

Tr Tr Tr ND

13.02 7.77 7.59 7.78

(94)a (86)b (86)b (88)b

(12)a (7)ab (8)ab (3)b

a

Values in parentheses are the proportion (%) of the total manure phosphorus determined by microwave digestion. Values in parentheses are the proportion (%) of the NaOH–EDTA extracted phosphorus. c Values for phosphate monoesters include myo-inositol hexakisphosphate and other monoesters. Tr, trace; ND, not detected. b

Feeding microbial phytase as a supplement There are several different types of phytase enzymes (see Mullaney and Ullah, Chapter 7, this volume), although they all catalyse the release of phosphate residues from myo-inositol hexakisphosphate. Phytase supplements are now a common component of animal diets and have been successful in reducing phosphorus concentrations in manures (see Lei and Porres, Chapter 9, this volume). However, the effects on manure phosphorus composition and therefore manure phosphorus behaviour in soils are poorly understood. It would be expected that manures from diets that included phytase would have less myoinositol hexakisphosphate than equivalent diets without phytase. This was the case in a study of manures from swine fed diets with and without phytase (Baxter et al., 2003). Concentrations of myo-inositol hexakisphosphate in fresh swine manure were decreased by 2.0–3.9 g P/kg by

adding phytase to the feed. However, during storage of manure from the normal diet for 150 days, myo-inositol hexakisphosphate as a percentage of total phosphorus decreased from 15.5% to 8.5%, which was attributed to microbial degradation. For the phytase-amended diet the decrease in myo-inositol hexakisphosphate during storage was only between 9.1% and 9.8%, indicating hydrolysis by the added phytase prior to excretion (Baxter et al., 2003). Therefore, after 150 days of storage, there was no significant difference in myo-inositol hexakisphosphate concentrations in swine manures from the two diets. Maguire et al. (2004) grew three flocks of broilers and two flocks of turkeys on the same bed of litter using diets that were ‘high’ and ‘low’ in non-phytate phosphorus with and without phytase additions. Concentrations of myo-inositol hexakisphosphate in both broiler and turkey litters from diets that included phytase were consistently lower than in litters from equivalent non-phytase diets (Table 10.4). Inorganic phosphate levels in the broiler and turkey litters

Table 10.4. Dietary studies where phytase has been used to reduce the total phosphorus concentrations in poultry manures and the influence on manure phytate content. Means followed by the same letter (within each column and for each study) are not significantly different (P > 0.05). Manure characteristics (g P/kg dry wt) Diet, non-phytate P (%)

Phytase addition

Total P

WSPa

Phytate

WSP/total P ratio

Phytate P/ total P ratio

Turkey Turkey Turkey Turkey

0.56 0.48 0.42 0.34

No Yes No Yes

17.8a 13.5b 11.8c 11.0d

6.4a 6.3a 5.1b 5.0b

5.09 3.45 4.89 3.65

0.36 0.47 0.43 0.45

0.28 0.26 0.41 0.33

Maguire et al. (2004)

Broiler Broiler Broiler Broiler

0.36 0.26 0.29 0.20

No Yes No Yes

14.1a 10.8c 11.7b 9.7d

4.7a 4.2a 2.2b 2.6b

7.83 4.88 7.32 5.35

0.33 0.39 0.19 0.27

0.56 0.45 0.63 0.55

Maguire et al. (2004)

Broiler Broiler Broiler Broiler

0.36 0.26 0.29 0.23

No Yes No Yes

13.6a 10.7bc 11.2b 9.6c

1.1a 1.0a 0.9a 0.6a

7.8 5.4 7.3 4.9

0.08 0.09 0.08 0.06

0.57 0.50 0.65 0.50

McGrath et al. (2005)

References

Environmental Implications in Animal Manures

Animal

a

WSP = water-soluble phosphate in manure.

157

158

A.B. Leytem and R.O. Maguire

were largely unaffected by dietary phytase. This was most likely due to the benefit of decreased dietary inorganic phosphate supplements being cancelled out by the increased phytate hydrolysis by dietary phytase. McGrath et al. (2005) determined myo-inositol hexakisphosphate in litters from broilers fed a variety of diets with and without phytase addition, and found that concentrations were lower in litter from diets containing phytase than from diets without phytase (Table 10.4). Toor et al. (2005) analysed turkey manure and broiler litter samples from diets with and without phytase using X-ray absorption near-edge structure spectroscopy. Although detection of organic phosphates was difficult using this technique, the authors concluded that dietary phytase addition decreased myo-inositol hexakisphosphate concentrations in manures and litters, and that dicalcium phosphate was the most abundant form of phosphorus present. There has been some discussion in the literature as to whether residual dietary phytase will continue to hydrolyse myo-inositol hexakisphosphate in manures following excretion, hence making phosphorus more water-soluble. Angel et al. (2005) used combinations of boiling poultry and swine manures, or added antibiotics, to show that dietary phytase supplementation had no effect on phytate hydrolysis following excretion. These authors concluded that the ‘increase in water-soluble phosphorus as a percent of total phosphorus post excretion is a function of excreta microbial activity and not dietary phytase addition’ (Angel et al., 2005). McGrath et al. (2005) stored broiler litters generated from diets ‘high’ and ‘low’ in phosphorus, with and without phytase, at two different moisture contents for 440 days. By comparing the interactions of storage time and moisture, they showed that myo-inositol hexakisphosphate concentrations decreased through time only in litter that was stored ‘wet’. This was unrelated to dietary phytase and was instead attributed to enhanced microbial activity in the wet litter (McGrath et al., 2005). Maguire et al. (2006) fed broiler breeders diets ‘high’ and ‘low’ in dietary non-phytate phosphorus, with and without phytase. Soluble phosphorus was similar in manure from under the feeder as in a clean area, indicating no effect of spilled feed whether or not it included phytase. However, under the drinker, manure moisture and soluble

phosphorus were higher irrespective of the diet, presumably due to increased microbial activity breaking down myo-inositol hexakisphosphate into more soluble forms. The effects of manurederived phytase in soils are unknown, although discussion of the interactions of phytase with soil constituents can be found elsewhere in this volume (see George et al., Chapter 14).

Combining low-phytate grains and phytase In addition to research on low-phytate grains or phytase alone, a few studies have investigated a combination of low-phytate grains and phytase. Baxter et al. (2003) reported that such a combination decreased myo-inositol hexakisphosphate in fresh swine manures more than either approach individually (Table 10.5). This trend was also seen in broiler litters, in which myo-inositol hexakisphosphate decreased from 20% of total phosphorus in a normal maize diet to 12% and 10% in diets containing low-phytate maize and lowphytate maize plus phytase, respectively (Toor et al., 2005; Table 10.5). Other studies combined phytase and low-phytate grains in poultry diets and reported reductions of 27–45% of total phosphorus and 27–49% of water-extractable phosphate in the litter, although none determined myo-inositol hexakisphosphate directly (Applegate et al., 2003; Miles et al., 2003; Penn et al., 2004).

Manure phosphorus composition and phosphorus solubility in soil Manipulating the diets of monogastric animals can have a large impact on the amount of myoinositol hexakisphosphate excreted from swine, poultry and fish. In addition, storage of manure prior to land application can also influence inositol phosphate concentrations by promoting microbial degradation. This raises an important question: Do differences in inositol phosphate concentrations influence the solubility and potential transport of manure phosphorus to water bodies following application to soil? Release of soluble phosphorus from manureamended soil varies considerably depending on the source of the manure applied (i.e. animal

Environmental Implications in Animal Manures

159

Table 10.5. Dietary studies utilizing low-phytate grains with and without the addition of phytase and the effect on manure phytate content. Means followed by the same letter (within column for each study) are not significantly different at P = 0.05.

Total P

WSPa

WSP/total P Phytate/total P ratio ratio

Animal

Diet

Phytase

Broiler

Normal maize

No

22.4a

12.6

0.56

20

Broiler

Low-phytate maize Low-phytate maize

No

20.1b

13.5

0.67

12

Yes

15.7c

12.0

0.76

10

Swine

Normal maize

No

25.5a

11.9a

0.47

15

Swine

Low-phytate maize Low-phytate maize

No

20.7b

10.8a

0.52

8

Yes

15.2c

7.9b

0.52

5

Broiler

Swine

g P/kg dry weight

Reference Toor et al. (2005)

Baxter et al. (2003)

a

WSP = water-soluble phosphate in manure.

species, diets fed, manure handling and storage). This is primarily due to differences in the concentrations of total and soluble phosphorus in the manure (Sharpley and Moyer, 2000; Kleinman et al., 2002a,b; Vadas et al., 2004), but may also be due in part to variability in other physical and chemical properties of the manure. Inorganic phosphate is relatively soluble in soils compared to myo-inositol hexakisphosphate, which is strongly retained and unlikely to be lost as soluble phosphorus in runoff (Anderson et al., 1974; Leytem et al., 2002). Therefore, variability of the phosphorus composition of manures, either due to differences in species, manure-handling techniques or through dietary manipulation, could increase phosphorus transport from land-applied manures to water bodies (Vadas et al., 2004). When a variety of manures (swine, dairy and beef cattle manures that were handled/stored differently) were incorporated into semiarid calcareous soils, there was no significant correlation between myo-inositol hexakisphosphate content (ranging between 0% and 8% of total phosphorus) and soil phosphorus solubility (Leytem and Westermann, 2005; Fig. 10.2a). In this instance, the small amounts of myo-inositol hexakisphosphate in the manures were probably insufficient to influence phosphorus solubility in the soil. Instead, phosphorus solubility was clearly influ-

enced by the amount of carbon added to the soil (Fig. 10.2b). When poultry manures were added to a similar calcareous soil, the amount of myo-inositol hexakisphosphate in the manures, which ranged between 35% and 80% of total phosphorus, was strongly and negatively correlated with bicarbonate-extractable soil phosphate, following manure application (Fig. 10.3a). Manures were applied at the same total phosphorus rate, so this correlation was almost certainly due to the greater proportion of water-soluble phosphate added in manure with lower myo-inositol hexakisphosphate concentrations. However, the relationship was transient, becoming insignificant after 9 weeks of incubation (Fig. 10.3b). This demonstrates clearly that when manures are applied on the basis of phosphorus content, the proportion of myo-inositol hexakisphosphate, and therefore of water-soluble phosphate, has a strong influence on the solubility of the manure phosphorus soon after application. Extractable phosphate concentrations increased between the second and ninth week of incubation and were correlated with the amount of myo-inositol hexakisphosphate in the manures. In other words, manures with more myo-inositol hexakisphosphate caused greater increases in extractable soil phosphate over time. Analysis of

160

A.B. Leytem and R.O. Maguire

(a)

(b)

Bicarbonate-extractable P (mg P/kg)

40 r 2 = 0.16; P = 0.43

r 2 = 0.81; P = 0.01

30

20

10

0 0

2

4

6

8

0

20

40

60

80

100

120

Manure phytate (% total P) Manure carbon/phosphorus ratio Fig. 10.2. Relationship between bicarbonate-extractable phosphate and (a) manure phytate concentration and (b) manure carbon/phosphorus ratio for six manures of varying origin added to a calcareous arable soil (Portneuf silt loam) from Idaho, USA, containing 0.75% organic carbon, pH 7.6 and 18% clay. (From Leytem and Westermann, 2005.)

the manure-amended soils immediately following incorporation (Fig. 10.4a) and after 9 weeks of incubation (Fig. 10.4b) using solution 31P NMR spectroscopy demonstrated the hydrolysis of myoinositol hexakisphosphate in the soil, strongly

Bicarbonate-extractable P (mg P/kg)

(a)

suggesting that this was responsible for the increase in extractable phosphate. Although myo-inositol hexakisphosphate is strongly bound in soils, microbes in the semiarid calcareous soil were able to break it down into (b)

12

8

4 2 weeks

9 weeks

r 2 = 0.99; P = 0.002

r 2 = 0.48; P = 0.193

0 0

20

40

60

80

100 0

20

40

60

80

100

Manure phytate (% total P) Fig. 10.3. Relationship between the phytate concentration in poultry manure and the bicarbonateextractable phosphate in manure-amended soil following (a) 2 weeks of incubation and (b) 9 weeks of incubation. The soil was a calcareous arable soil (Portneuf silt loam) from Idaho, USA, containing 0.75% organic carbon, pH 7.6 and 18% clay. (From Leytem et al., 2006.)

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(a) 6.3 5.1 4.7 4.5

5.9

7.0 6.5 6.0 5.5 5.0 4.5 4.0

(b)

7.0 6.5 6.0 5.5 5.0 4.5 4.0

6

4

2

0

Chemical shift (ppm) Fig. 10.4. Solution 31P nuclear magnetic resonance (NMR) spectra of extracts of a soil amended with poultry manure (a) immediately following incorporation and (b) after 9 weeks of incubation. The peak at 6.3 ppm is inorganic phosphate, while the other four labelled signals are from myo-inositol hexakisphosphate. The spectra demonstrate the relatively rapid hydrolysis of manure-derived myoinositol hexakisphosphate in soil. (From Leytem et al., 2006.)

inorganic phosphate within a few weeks. It would therefore not be expected to accumulate in these soils following successive manure applications. This confirms the evidence for the relative bioavailability of inositol phosphates in calcareous soils (Turner et al., 2003) and may explain why some contain no detectable phytate (see Turner, Chapter 12, this volume). In contrast, the same manures applied to an acidic soil showed no correlation between added manure myo-inositol hexakisphosphate and extractable soil phosphate (Mehlich-3 extraction) on any of the sampling dates, with only the manure carbon/phosphorus ratio being correlated to the extractable phosphate concentrations (r2 = 0.84 at 2 weeks of incubation; data not shown). The solubility of phosphorus in manureamended soils seems to be influenced by the characteristics of the manure applied. In the

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short term, manures with large concentrations of myo-inositol hexakisphosphate can demonstrate lower phosphorus solubility on calcareous soils, although this trend does not seem to hold true for acidic soils. However, due to microbial breakdown of myo-inositol hexakisphosphate in applied manures and concurrent release of soluble phosphate, these differences are likely to become insignificant over time. Other manure properties, particularly the carbon content, seem to exert a large influence on phosphorus solubility following application to both calcareous and acidic soils (Leytem et al., 2005), presumably due to stimulation of the microbial biomass and fixation of phosphorus in microbial tissue. This means that the addition of manure results in a lower soluble phosphorus concentration than would be expected from mineral phosphate fertilizer application. It therefore follows that in the long term the most important factor to consider for land application of manures is total phosphorus, rather than the form of the phosphorus applied. An important impact of manure inositol phosphates on the loss of phosphorus to water bodies involves erosion and transport of particulate phosphorus. Erosion can be severe on agricultural land and is potentially responsible for the movement of large amounts of inositol phosphates to water bodies (see McKelvie, Chapter 16, this volume). Erosion can be promoted by inositol phosphates in manures due to the dispersion of soil colloids following sorption to soil components (see Celi and Barberis, Chapter 13, this volume). There is almost no information on inositol phosphate transport in particulate material from agricultural land, and it is not discussed further here. However, several stereoisomeric forms of inositol hexakisphosphate have been reported from riverine-suspended solids (Suzumura and Kamatani, 1995). More information can be found in a detailed review of organic phosphorus transfer from soils to water bodies (Turner, 2005).

Dietary Manipulation and the Environmental Fate of Manure Phosphorus Manures from low-phytate feed Although the total phosphorus excreted from monogastric animals fed a variety of low-phytate

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grains has been shown to be significantly reduced, the impacts of these manures on potential phosphorus losses following long-term application to agricultural land have not been studied. One of the primary reasons for this is the lack of sufficient quantities of manure needed for fieldscale assessments, particularly multi-year projects. Investigation is therefore limited to laboratoryscale studies. Gollany et al. (2003) showed a 10% reduction in manure phosphorus availability when manure from swine fed low-phytate maize-based diets vs. normal maize diets was incorporated into a silt loam soil. Leytem et al. (2005) incorporated manure from swine fed a variety of lowphytate barley-based diets and found no significant relationship between the amount of myo-inositol hexakisphosphate added in the manures and bicarbonate-extractable phosphate in soil (Fig. 10.5a). However, as with previous studies, there was a strong relationship between the amounts of carbon added with the manures and the bicarbonate-extractable phosphate (Fig. 10.5b). As the amount of phosphorus excreted by animals fed low-phytate grains is reduced, there is a corresponding increase in the manure carbon/ phosphorus ratio, which can enhance the stabilization of phosphorus in manure-amended soils compared with soils amended with manures from normal grain-based diets. Therefore, even when

applied on the same total phosphorus basis, there is a potential environmental benefit to feeding low-phytate grains when the subsequent manures are land-applied, at least in the short term.

Manures from phytase-amended feed Studies have consistently shown reductions in manure total phosphorus and myo-inositol hexakisphosphate from swine and poultry that have been fed diets with phytase, but only when, as recommended, inorganic phosphate supplementation is reduced to account for enhanced phosphorus availability due to phytase addition. However, there has been some disagreement over the effect of added phytase on manure water-extractable phosphate, which is important because it is linked directly to phosphorus losses in runoff (Maguire et al., 2005a). Dietary phytase addition can decrease total manure phosphorus concentrations by as much as 45% for poultry and 40% for swine (see Lei and Porres, Chapter 9, this volume). These reductions are important, as total phosphorus determines build-up or decline in soil test phosphorus following land application of manures. This is particularly true where manure is applied on the basis of nitrogen content – the effects of changes in manure

Bicarbonate-extractable P (mg P/kg)

(a)

(b)

100 r 2 < 0.01; P = 0.51

r 2 = 0.80; P < 0.001

80

60

40

20 0

2

4

6

8

10

20

40

60

80

Manure phytate (% total P) Manure carbon/phosphorus ratio Fig. 10.5. The relationship between bicarbonate-extractable phosphate and (a) manure phytate concentration or (b) manure carbon/phosphorus ratio for manures from swine fed low-phytate grain diets applied to a calcareous arable soil (Portneuf silt loam) from Idaho, USA, containing 0.75% organic carbon, pH 7.6 and 18% clay. (From Leytem et al., 2005.)

Environmental Implications in Animal Manures

phosphorus composition are therefore only likely to become relevant when manure is applied on the basis of phosphorus content. Several studies have surface-applied manures and litters derived from phytase-amended diets and measured phosphorus in runoff. Smith et al. (2004a) reported that although dietary phytase additions decreased the water-extractable phosphate in swine manure, this had no significant effect on soluble phosphorus losses in runoff from manured soils, relative to manure from a nonphytase-amended diet. This was surprising because equivalent weights of manures were applied, so manures with smaller concentrations of water-soluble phosphate (i.e. from phytaseamended diets) were expected to yield less soluble phosphate in runoff. In a similar study, however, soluble phosphate concentrations in runoff immediately following the application of poultry litter from a phytase-amended diet were lower than from soils that received litter from a normal diet (Smith et al., 2004b). Again, manure was applied on a weight basis and, importantly, the effect became insignificant when three consecutive rainfall events were included. It should be noted that in both studies the application of alum (aluminium sulphate) to the litters considerably reduced soluble phosphate in litter and in runoff following litter application to soil. In one study in which dietary phytase significantly increased manure water-extractable phosphate, Vadas et al. (2004) reported no significant differences in soluble phosphate concentrations in runoff between soils amended with poultry manures from phytase and non-phytase-amended diets, even when manures were applied at the same total phosphorus rate. Using turkey and broiler litters from equivalent phytase- and non-phytase-amended diets, Maguire et al. (2004, 2005b) found that dietary phytase decreased myo-inositol hexakisphosphate in litters, but generally had little effect on manure inorganic phosphate or soluble phosphate losses in runoff when manures were incorporated into soil prior to rainfall. This occurred whether litter was applied on the basis of nitrogen or phosphorus content. Where more than one runoff event was conducted, soluble phosphate losses decreased as the number of runoff events increased, and the effects of diet and manure characteristics became less significant. These data highlight the point that the soluble phosphorus in

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manure has a greater impact on runoff soluble phosphate concentrations in the short term than in the long term (Penn et al., 2004; Smith et al., 2004b; Maguire et al., 2005b). However, we still must consider the fact that long-term land application of manures results in the accumulation of a large pool of phosphorus, which may be available for release to runoff water over time. The reduction in total manure phosphorus with phytase additions has the long-term benefit of reducing total phosphorus additions to fields receiving continual nitrogen-based manure applications that overapply phosphorus compared to crop needs.

Manures from low-phytate grains and phytase-amended feeds As already discussed, combining low-phytate grains and phytase was shown to result in greater reductions in manure total phosphorus than either strategy on its own. It has also been shown to reduce water-extractable phosphate by 27–49% (Maguire et al., 2005a). Smith et al. (2004b) reported that adding phytase to poultry diets containing low-phytate maize led to less soluble phosphate in runoff compared to that from a normal diet, but was not different to soluble phosphate in runoff from diets containing phytase or low-phytate maize on their own when manures were surface-applied at the same total phosphorus rate. Penn et al. (2004) observed similar concentrations of soluble phosphate in runoff from soils receiving surface application of turkey manure (same total phosphorus applied) from normal or low-phytate maize plus phytase diets. As there are only a limited number of studies measuring runoff from soils amended with these manures, it is too early to draw firm conclusions. However, the consistent reduction in total phosphorus and waterextractable phosphate in the manures suggests a clear benefit in terms of water quality.

Summary Research to date has shown manure composition to be heavily dependent on both animal species and diet. In particular, differences in feed composition and phytase supplementation mean that

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manures from monogastric animals contain a wide range of myo-inositol hexakisphosphate concentrations. However, manure tends to be stored for long periods of time prior to land application, which allows microbial activity to break down a large fraction of the myo-inositol hexakisphosphate. This creates manures that have low myoinositol hexakisphosphate concentrations when they are eventually land-applied. An important consequence is that other manure characteristics, such as the carbon/phosphorus ratio, may have a greater influence on subsequent phosphorus solubility in the short term than the phosphorus composition of the manure upon excretion from the animal. This must be considered when assessing the effects of dietary manipulation on the environmental impact of manure phosphorus. When manure is applied to soil, a variety of factors can influence the phosphorus solubility and the potential for phosphorus transport to water bodies. In the case of surface-applied manure, the water-extractable phosphate concentration has the greatest influence on soluble phosphate losses when rainfall immediately follows manure application. When manures are incorporated into soils, other factors control phosphorus solubility and the potential for phosphorus losses to water bodies. In calcareous soils with low organic matter contents, phosphorus sorption can be influenced in the short term by the myo-inositol hexakisphosphate content of the manure, because manure with large concentrations of myo-inositol hexakisphosphate lead to small increases in soil phosphate solubility compared with manures dominated by inorganic phosphate. However, this effect is reduced as myo-inositol hexakisphosphate undergoes hydrolysis and contributes to the extractable phosphate pool, at which point other factors, such as the manure carbon/phosphorus ratio, determine differences in phosphate solubility. In contrast, when manures are applied to acidic soils, there seems to be no influence of myoinositol hexakisphosphate content on extractable soil phosphate, and other manure characteristics may have a greater influence on phosphorus solubility. In situations where phosphorus losses are dominated by soil erosion and particulate phosphorus losses, the phosphorus concentration in the soil will overwhelm any influence of the applied manure phosphorus forms. Concern has been expressed about the potential negative environmental implications of

diet alteration on phosphorus losses from manureamended soils, but given the urgent requirement to reduce total phosphorus concentrations in manures in areas of high livestock density, dietary manipulation is overwhelmingly beneficial. Such manipulation may increase the proportion of the manure phosphorus that is soluble in water, but this is likely to have negative environmental consequences only when manure is applied on a phosphorus basis and without prolonged storage prior to land application. If manures are applied on an equivalent weight or nitrogen basis, diet modification will result in less total phosphorus being added to soils and therefore a reduction in soil test phosphorus build-up over time. This in turn decreases the risk of phosphorus transfer to water bodies. In addition, most research indicates a reduction or no increase in phosphorus losses in runoff from soils amended with manures from modified diets compared with normal diets, when these are applied on an equivalent phosphorus basis (surface application or incorporation of manures). It therefore seems likely that in most cases there is no enhanced environmental risk from dietary modification and associated changes in manure phosphorus composition.

Future Research Needs There is an increasing body of research aimed at understanding the influence of manure phosphorus composition on the potential environmental impacts related to land application of manure. At present, few studies have determined manure phosphorus composition using techniques such as solution 31P NMR spectroscopy, yet this information provides valuable insight into the behaviour of phosphorus in manure after land application and can help identify the potential risks of modifying manures through diet manipulation. The study of dietary impacts on manure phosphorus composition and subsequent environmental risk is becoming more important. There are few studies that have detailed the impacts of altering animal diets on manure phosphorus composition, and these have focused primarily on phosphorus in feeds (i.e. non-phytate phosphorus levels and the use of phytase). Dietary components, such as the calcium/phosphorus ratio in feeds, micronutrient additions and carbon

Environmental Implications in Animal Manures

composition of feeds, can alter the manure phosphorus composition and influence waterextractable phosphate, but have not been investigated in detail (see also Dao, Chapter 11, this volume). There is evidence that calcium and other divalent cations often found in micronutrient supplements can bind with myo-inositol hexakisphosphate, making both less available during digestion (Maenz et al., 1999). Future studies should therefore look beyond just dietary phosphorus in order to understand the extent to which we can alter the phosphorus composition in manures and maximize the benefits of dietary manipulation. The use of low-phytate grains in animal feeding operations has received considerable interest (see Raboy, Chapter 8, this volume) and further research will be necessary as new grains become available, especially as these become economically viable. Low-phytate grains have an advantage over phytase addition, because they

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minimize the interference that dietary inputs (such as calcium and other micronutrients) may have on phytate digestion and phytase efficacy. An important drawback at this point to using low-phytate grains is the issue of identity preservation (ability to keep low-phytate grains separate from other grains during processing), which will hopefully be overcome in the future. Now that modified diets (phytase additions, low-phytate grains and lower phosphorus) are widely implemented, there is a need for long-term studies to determine the environmental effects of manure application resulting from these diets and the effects on soil phosphorus forms. There are no long-term trials studying the effect of land-applied manures from low-phytate diets on soil organic matter, soil phosphorus availability and forms, or phosphorus losses in runoff. Given the importance of understanding the impact of intensive animal operations on the phosphorus pollution of water bodies, such studies are urgently required.

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Maguire, R.O., Plumstead, P.W. and Brake, J. (2006) Impact of diet, moisture, location and storage on soluble phosphorus in broiler breeder manure. Journal of Environmental Quality 35, 858–865. McAuliffe, C. and Peech, M. (1949) Utilization by plants of phosphorus in farm manure: I. Soil Science 68, 185–195. McCuaig, L.W., Davies, M.I. and Motzok, I. (1972) Intestinal alkaline phosphatase and phytase of chicks: effects of dietary magnesium, calcium, phosphorus and thyroactive casein. Poultry Science 51, 526–530. McGrath, J.M., Sims, J.T., Maguire, R.O., Saylor, W.W., Angel, R. and Turner, B.L. (2005) Broiler diet modification and litter storage: impacts on phosphorus in litters, soils, and runoff. Journal of Environmental Quality 34, 1896–1909. Mikkelsen, R.L. (2000) Beneficial use of swine by-products: opportunities for the future. In: Powers, J.F. and Dick, W.A. (eds) Land Application of Agricultural, Industrial, and Municipal By-products. Soil Science Society of America, Madison, Wisconsin, pp. 451–480. Miles, D.M., Moore, P.A., Smith, D.R., Rice, D.W., Stilborn, H.L., Rowe, D.R., Lott, B.D., Branton, S.L. and Simmons, J.D. (2003) Total and water-soluble phosphorus in broiler litter over three flocks with alum litter treatment and dietary inclusion of high available phosphorus corn and phytase supplementation. Poultry Science 82, 1544–1549. Mullins, G., Joern, B. and Moore, P. (2005) By-product phosphorus: sources, characteristics and management. In: Sims, J.T. and Sharpley, A.N. (eds) Phosphorus: Agriculture and the Envrionment. Agronomy Monograph No. 46. American Society of Agronomy, Madison, Wisconsin, pp. 829–879. Penn, C.J., Mullins, G.L., Zelazny, L.W., Warren, J.G. and McGrath, J.M. (2004) Surface runoff losses of phosphorus from Virginia soils amended with turkey manure using phytase and high available phosphorus corn diets. Journal of Environmental Quality 33, 1431–1439. Peperzak, P., Caldwell, A.G., Hunziker, R.R. and Black, C.A. (1959) Phosphorus fractions in manures. Soil Science 87, 293–302. Raboy, V., Gerbasi, P.F., Young, K.A., Stoneberg, S.D., Pickett, S.G., Bauman, A.T., Murthy, P.P.N., Sheridan, W.F. and Ertl, D.S. (2000) Origin and seed phenotype of maize low phytic acid 1–1 and low phytic acid 2–1. Plant Physiology 124, 355–368. Rather, J.B. (1918) The Utilization of Phytin Phosphorus by the Pig. Bulletin No. 147. University of Arkansas College of Agriculture, Agriculture Experiment Station, Fayetteville, Arkansas. Sharpley, A.N. (1996) Availability of residual phosphorus in manured soils. Soil Science Society of America Journal 60, 1583–1588. Sharpley, A.N. and Moyer, B. (2000) Phosphorus forms in manure and compost and their release during simulated rainfall. Journal of Environmental Quality 19, 1462–1469. Sharpley, A.N., Chapra, S.C., Wedephol, R., Sims, J.T., Daniel, T.C. and Reddy, K.R. (1994) Managing agricultural phosphorus for protection of surface waters: issues and options. Journal of Environmental Quality 23, 437–451. Sharpley, A., Gburek, W. and Heathwaite, L. (1998) Agricultural phosphorus and water quality: sources, transport and management. Agriculture and Food Science of Finland 7, 297–314. Sims, J.T., Simard, R.R. and Joern, B.C. (1998) Phosphorus loss in agricultural drainage: historical perspective and current research. Journal of Environmental Quality 27, 277–293. Sims, J.T., Edwards, A.C., Schoumans, O.F. and Simard, R.R. (2000) Integrating soil phosphorus testing into environmentally based agricultural management practices. Journal of Environmental Quality 29, 60–71. Smith, D.R., Moore, P.A. Jr, Maxwell, C.V., Haggard, B.E. and Daniel, T.C. (2004a) Reducing phosphorus runoff from swine manure with dietary phytase and aluminum chloride. Journal of Environmental Quality 33, 1048–1054. Smith, D.R., Moore, P.A. Jr, Miles, D.M., Haggard, B.E. and Daniel, T.C. (2004b) Decreasing phosphorus runoff from land applied poultry litter with dietary modifications and alum addition. Journal of Environmental Quality 33, 2210–2216. Spencer, J.D., Allee, G.L. and Sauber, T.E. (2000) Phosphorus bioavailability and digestibility of normal and genetically modified low-phytate corn for pigs. Journal of Animal Science 78, 675–681. Suzumura, M. and Kamatani, A. (1995) Origin and distribution of inositol hexaphosphate in estuarine and coastal sediments. Limnology and Oceanography 40, 1254–1261. Toor, G.S., Peak, J.D. and Sims, J.T. (2005) Phosphorus speciation in broiler litter and turkey manure produced from modified diets. Journal of Environmental Quality 34, 687–697. Turner, B.L. (2004) Optimizing phosphorus characterization in animal manures by phosphorus-31 nuclear magnetic resonance spectroscopy. Journal of Environmental Quality 33, 757–766. Turner, B.L. (2005) Organic phosphorus transfer from terrestrial to aquatic environments. In: Turner, B.L., Frossard, E. and Baldwin, D.S. (eds) Organic Phosphorus in the Environment. CAB International, Wallingford, UK, pp. 269–294.

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Turner, B.L. and Leytem, A.B. (2004) Phosphorus compounds in sequential extracts of animal manures: chemical speciation and a novel fractionation procedure. Environmental Science and Technology 38, 6101–6108. Turner, B.L., Papházy, M., Haygarth, P.M. and McKelvie, I.D. (2002) Inositol phosphates in the environment. Philosophical Transactions of the Royal Society, London, Series B 357, 449–469. Turner, B.L., Cade-Menun, B. J. and Westermann, D.T. (2003) Organic phosphorus composition and potential bioavailability in semi-arid arable soils of the western United States. Soil Science Society of America Journal 67, 1168–1179. US Geological Survey (1999). The quality of our nation’s waters: nutrients and pesticides. USGS Information Services, Denver, Colorado. Vadas, P.A., Meisinger, J. J., Sikora, L.J., McMurtry, J.P. and Sefton, A.E. (2004) Effect of poultry diet on phosphorus in runoff from soils amended with poultry manure and compost. Journal of Environmental Quality 33, 1845–1854. Veum, T.L., Ledoux, D.R., Bollinger, D.W., Raboy, V. and Cook, A. (2002) Low-phytic acid barley improves calcium and phosphorus utilization and growth performance in growing pigs. Journal of Animal Science 80, 2663–2670. Weinhold, B. J. and Miller, P.S. (2004) Phosphorus fractions in manure from swine fed traditional and low-phytate corn diets. Journal of Environmental Quality 33, 389–393.

11

Ligand Effects on Inositol Phosphate Solubility and Bioavailability in Animal Manures Thanh H. Dao

United States Department of Agriculture–Agricultural Research Service, Beltsville Agricultural Research Center, Room 121, 10300 Baltimore Avenue, Building 306 BARC-EAST, Beltsville, MD 20705, USA

In regions with high concentration of confined animal production operations, continuous manure application over many years has resulted in agricultural soils that contain high levels of phosphorus (Simard et al., 1995; Frossard et al., 2000; Pautler and Sims, 2000; Zhang et al., 2002; Lehmann et al., 2005). The build-up increases the potential for phosphorus leaching and transport in runoff to nearby water bodies, with the associated risk of water quality impairment (Gerritse and Eksteen, 1978; Chardon et al., 1997; Koopmans et al., 2003). A large proportion of the phosphorus in animal manures can be in the form of inositol phosphates, which occur mainly as salts of myo-inositol hexakisphosphate (phytate) (Peperzak et al., 1959; Gerritse and Eksteen, 1978; Dao, 2004b; Toor et al., 2005a). As a result, there is considerable interest in the identification and behaviour of inositol phosphates in manures and soils. Other chapters in this volume discuss inositol phosphates in animal nutrition, excretion in manures and the fate of manure phosphorus in the environment (see Lei and Porres, Chapter 9, and Leytem and Maguire, Chapter 10, this volume). This chapter examines the processes that influence the solubilization and dephosphorylation of these inositol phosphates in manures. This is of particular importance in an environmental context, because these processes influence the release and bioavailability of inositol phosphates in soils and water bodies. An in situ ligand-based

enzymatic hydrolysis method has been developed to assess the bioavailability of inositol phosphates in manure. The procedure illustrates that characterization of phosphorus in manures, based on a combination of chemical and biological assays, may more appropriately reflect the availability of inositol phosphates to organisms and plants. The mild fractionation approach may also reveal important information about the stability and biological activity of complexed forms of inositol phosphates, which are often considered to be chemically unreactive in the environment (see McKelvie, Chapter 16, this volume).

Assessing the Solubility and Release of Inositol Phosphates in Animal Manures Mixed salts of myo-inositol hexakisphosphate account for 60–90% of the total phosphorus in seeds of wheat (Triticum aestivum L.), rice (Oryza sativa L.), maize (Zea mays L.), soybean [Glycine max (L.) Merr.] and mung bean [Vigna radiata (L.) R. Wilczek var. radiata] (Scott and Loewus, 1986; Lott et al., 2000; see Raboy, Chapter 8, this volume). myo-Inositol hexakisphosphate is also found in leaves of a variety of plants, including Arabidopsis thaliana (L.) Heynh. (Bentsink et al., 2003), Telfairia occidentalis Hook F. (Ladeji et al.,

©CAB International 2007. Inositol Phosphates: Linking Agriculture and the Environment (eds B.L. Turner, A.E. Richardson and E.J. Mullaney)

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1995), Euphorbia hirta L. and Launaea taraxacifolia (Wild.) Amin ex C. Jeffrey (Wallace et al., 1998). Diets of domestic livestock and poultry therefore contain abundant myo-inositol hexakisphosphate. However, swine (Sus scrofa domesticus) and poultry (Gallus gallus domesticus) are monogastric and do not possess sufficient levels of phytase enzymes in their digestive system to break down inositol hexakisphosphate, which therefore occurs in large concentrations in their manures (Zyla et al., 2000; Council for Agricultural Science and Technology, 2002; Turner, 2004; Toor et al., 2005b; see Leytem and Maguire, Chapter 10, this volume). In contrast, it is generally thought that in ruminant livestock such as cattle (Bos taurus), sheep (Ovis aries) and goats (Capra aegagrus hircus) phytases produced by the rumen microflora and phytases in saliva and intestinal mucosa catalyse the hydrolysis of myo-inositol hexakisphosphate during the rumination process, releasing phosphate for assimilation by the animal. However, a number of recent studies have reported evidence for the presence of myo-inositol hexakisphosphate in manure from cattle fed grain-based diets (Dao, 2003; Jayasundera et al., 2005; Toor et al., 2005a). Inositol phosphates could escape into the manure of ruminants following reaction with polyvalent cations to form insoluble precipitates. However, the scarcity of information on the speciation of inositol phosphates in manure and their hydrolysis to release inorganic phosphate presents a major challenge to understanding phosphorus dynamics in manure and its fate in the environment following land application. To assess the composition and solubility of phosphorus in animal manure, some studies have adapted fractionation methods developed for soil phosphorus. Sequential fractionation procedures were developed to distinguish phosphorus pools based on the differential solubility of calcium, aluminium and iron phosphates in strong acids and bases (Chang and Jackson, 1957; Hedley et al., 1982; Kuo, 1996). Solidphase extraction media that included ionexchange resins or iron oxide-impregnated paper were used as anion sinks to measure the labile phosphorus fraction in soil (van Diest et al., 1960; Abrams and Jarrell, 1992; Chardon et al., 1996; Myers et al., 1999). However, methods designed for soils may not be readily applicable to manures. In particu-

lar, manures are composed of partly digested feed and are primarily organic, whereas soils have a mineral phase and an associated complex organic carbon phase. These matrices require caustic chemical treatments to release the different phosphorus forms associated with soil mineral and organic matter surfaces that exist as a range of insoluble inorganic and organic phosphate compounds in various stages of crystallization (Lindsay, 1979; Graf, 1986). The possibility that soil phosphorus analytical methods do not yield discrete chemical fractions also exists. Chemical transformations and transfers between phosphorus pools occur as a result of the choice of solvents, the chemical composition of the extractant solution or the harshness of the extraction conditions (Adams and Byrne, 1989; Leinweber et al., 1997; Turner, 2004; Turner and Leytem, 2004; McDowell and Stewart, 2005). For example, the concentration of sodium hydroxide in the extracting solution can alter the distribution of phosphate monoesters and diesters in alkaline extracts of animal manures and also influences spectral resolution in solution 31P nuclear magnetic resonance (NMR) spectroscopy (Leinweber et al., 1997; Turner, 2004). The extraction of phosphates by strong acidic and basic extractants is not limited to single species of phosphate; for example, aluminium phosphate and iron phosphate are both soluble in acidic extractants. Solubility products average 20.3 and 28 for aluminium and iron phosphates, and 32.5 and 96 for aluminium and iron hydroxy-phosphates, respectively (de Haas et al., 2001). Methods involving the enzymatic dephosphorylation of phosphorus-containing compounds have been used to characterize inositol phosphates and other groups of organic phosphorus in extracts of animal manures (He and Honeycutt, 2001; Dao, 2003) and soils (Shand and Smith, 1997; Hayes et al., 2000; Turner et al., 2002; Toor et al., 2003; Dao, 2004a; Dao et al., 2005). Phosphatases catalyse chemical reactions that release phosphate from various types of organic phosphorus compounds. That is, when a manure sample or an extract of a manure sample is incubated with a specific phosphatase, the release of inorganic phosphate in the reaction medium indicates the presence of a specific type of organic phosphorus compound and its concentration in the sample. For example, He and Honeycutt (2001) used a modified sequential extraction

Ligand Effects in Animal Manures

method for soil phosphorus (Hedley et al., 1982) and subjected the various extracts of swine and cattle manure to hydrolysis by acid and alkaline phosphatases, phytases, nuclease P1 and nucleotide pyrophosphatase. They found myo-inositol hexakisphosphate-like compounds and small quantities of phosphate diesters. This method appears to provide qualitative information on the composition and potential bioavailability of the manure organic phosphorus. However, enzymatic methods have been hampered by the low reactivity of hydrolysable organic phosphorus in extracts, which means that a large proportion of the organic phosphorus remains uncharacterized. For example, in the study described above (He and Honeycutt, 2001), hydrolysable phosphorus accounted for <50% of the organic phosphorus extracted by water and sodium hydroxide, and <15% of the organic phosphorus extracted by bicarbonate and hydrochloric acid. In addition, the extent of phosphate release from organic phosphorus substrates may also be affected by the specificity and purity of the various enzyme preparations. For example, commercial phytase preparations (e.g. phytase from Sigma Chemical Company, St Louis, Missouri, USA) express non-specific acid phosphatase activity, which can lead to an overestimation of phosphate release from inositol phosphates compared to the use of more highly purified and thus more specific phytases (Shand and Smith, 1997; Hayes et al., 2000). Ideally, enzyme-based assays to determine the phytase-hydrolysable phosphate should employ phytases with high specific activity for myo-inositol hexakisphosphate. The fact that relatively small proportions of organic phosphorus in extracts of manure and soil can be hydrolysed by phytase has been attributed to the complexation of organic phosphates with polyvalent cations. Dao (2003) observed that the inefficiency of enzymatic methods was not related to insufficient levels of enzyme activity or inositol phosphate substrates, but was due in part to the association of inositol phosphates with polyvalent cations that control their solubility and susceptibility to dephosphorylation. A similar situation was found in studies of in situ hydrolysis of phosphate monoesters in soils (Otani and Ae, 1999; Dao, 2004a). The phenomenon has important impacts on the solubility and reactivity of inositol phosphates in animal manures, and is explored later.

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Inositol Phosphate Reactivity with Polyvalent Cations The six phosphate moieties give myo-inositol hexakisphosphate the potential of 12 coordinate ligands for complexing cations. It complexes monovalent cations and stoichiometrically forms monomeric salts such as sodium phytate. Multivalent cations such as calcium or iron(III) ions can form intramolecular bonds, bridging two or more phosphate moieties on a single inositol molecule, resulting in monomeric inositol phosphates; intermolecular bonding can occur when two or more inositol phosphate molecules share a common multivalent cation and yield polymeric compounds (Vohra et al., 1965; Evans and Pierce, 1982; Pierce, 1985; Champagne et al., 1990). For example, chemical interactions between myoinositol hexakisphosphate and cations were evident in the finding that a high calcium/ phosphorus molar ratio in poultry diets interfered with the effectiveness of dietary phytases. The formation of insoluble calcium phytate under the intestinal conditions rendered the compound resistant to enzymatic hydrolysis (Qian et al., 1997; Kornegay, 2001). One would expect similar reactions involving lower-order inositol phosphates, although the chelation potential is reduced with a decrease in the number of phosphate moieties on the inositol molecule. For example, clinical studies showed that zinc and iron absorption in humans increased with myoinositol tris-, tetrakis- and pentakisphosphate, compared with the hexakisphosphate form (Sandström and Sandberg, 1992; Sandberg et al., 1999). Therefore, the extent to which polyvalent cations influence the physical state of inositol phosphates depends on three factors: the cation, the cation concentration or molar ratio of cation to inositol phosphate and the interaction with other cations present.

Effects of counterion valency and solution-phase pH By varying the molar ratio of cation to myoinositol hexakisphosphate, it was observed that dephosphorylation of myo-inositol hexakisphosphate was not affected by divalent calcium ions at a mole fraction of 1:6, in comparison to

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monovalent sodium ions (Fig. 11.1). Calcium phytate was soluble at low molar ratios of calcium to myo-inositol hexakisphosphate-phosphorus, but dephosphorylation was reduced by 50% when the molar ratio of calcium to myo-inositol hexakisphosphate increased to 6:1 (i.e. a 6:6 calcium/phosphate ratio), due to the precipitation of insoluble calcium phytate. Divalent calcium ions did not completely inhibit dephosphorylation to the same extent as aluminium and iron, because calcium phytate remains soluble at pH 4 (Wise and Gilburt, 1981). As a strong ligand, myo-inositol hexakisphosphate had a high affinity for aluminium and iron(III) ions, so dephosphorylation was progressively inhibited as the concentrations of these cations or the mole ratio of cation to myo-inositol hexakisphosphate increased (Fig. 11.1). In aluminium and iron treatments, phosphate release

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decreased by an average of 27% and 32% compared with the sodium treatment at 1:6 cation/ myo-inositol hexakisphosphate molar ratio or 0.25 mM of metal ions at pH 4.5. The phenomenon has been referred to as an ‘in solution’ sequestration of inositol phosphate; although no visible precipitate was observed, the formation of metal chelates sequestered and shielded some of the inositol phosphate from dephosphorylation by phytase (Dao, 2003). More than 80% and 99% inhibition of dephosphorylation was observed when molar ratios of aluminium or iron to myoinositol hexakisphosphate exceeded 3:6 and 6:6, respectively. myo-Inositol hexakisphosphate is more susceptible to dephosphorylation at pH 6 than pH 4.5 (Fig. 11.1b). The molecule has 12 ionizable protons and 6 of them have a pKa ≥ 5.2, while the remaining pKa values are <3.2 (Evans and Pierce, 1982).

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Fig. 11.1. Short-term kinetics of the dephosphorylation of myo-inositol hexakisphosphate as affected by polyvalent counterion concentrations and solution-phase pH (a=pH 4.5; b=pH 6.0). (Reprinted from Dao, 2003.)

Ligand Effects in Animal Manures

As solution pH increases, more metal hydroxide species, such as M(OH)2+, M(OH)2+ and M(OH)3 (where M = aluminium or iron) and consequently less Al(H2O)63+ or Fe(H2O)63+ aquo-metal ions, would exist in solution. The formation of more amorphous Al(OH)3 and Fe(OH)3 would reduce the effective concentrations of aluminium and iron that can react with myo-inositol hexakisphosphate. Also, more inositol phosphate remains in dissociated forms and is susceptible to dephosphorylation at pH 6.0, compared to pH 4.5. Therefore, the inhibition of dephosphorylation by aluminium and iron has been attributed primarily to chelation, resulting in sterically hindered forms of myo-inositol hexakisphosphate.

The presence and persistence of myo-inositol hexakisphosphate in ruminant excreta Similar reactions to those described above affected myo-inositol hexakisphosphate added to liquid dairy manure suspensions (Dao, 2003). Solutionphase phosphate increased with the addition of Aspergillus ficuum phytase, but this was reduced in samples with added polyvalent cations. The added phytases hydrolysed an average of 22 mmol P/kg dairy manure solids (native phytase activity in the manure was deactivated by steam sterilization). This demonstrated the presence of soluble, fully ionized and uncomplexed myo-inositol hexakisphosphate in dairy manure that was amenable to hydrolysis by added phytase. It did not represent all the inositol phosphates in dairy manure, although the phytase-hydrolysable phosphorus (PHP) assay was subsequently improved with the use of a combination of phytases and polydentate ligands (Dao, 2004a,b) (see below). On the basis of these results it was postulated that these sequestration reactions are mechanisms by which inositol phosphates in feed grain persist during passage through the animal, leading to excretion in the faeces of dairy cattle (Dao, 2003). In the animal digestive tract and in manure, myoinositol hexakisphosphate and lower-order inositol phosphates likely interact with polyvalent cations, primarily calcium, aluminium, iron and magnesium, and to a lesser extent with manganese, zinc and copper, as these micronutrients are added to the feeds as mineral supplements for nutritional

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and health considerations (National Research Council, 2001). The combination of the rapid rate of feed passage through high-productivity dairy cattle and the reduced susceptibility of complexed inositol phosphate to dephosphorylation by complexation with dietary polyvalent cations results in the persistence and excretion of complexed myoinositol hexakisphosphate in dairy manure. Thus, while rumen microflora or intestinal mucosa phytases are theoretically capable of hydrolysing myo-inositol hexakisphosphate in feed, this does not appear to happen in practical feeding regimens. Manure from dairy and beef cattle fed grain-based diets can therefore contain considerable amounts of myo-inositol hexakisphosphate ( Jayasundera et al., 2005; Toor et al., 2005a) and, as for monogastric animals, presents an environmental concern.

Characterizing the Relative Stability of Inorganic Phosphate and Inositol Phosphates in Animal Manure Metal chelate stability The reactivity of inositol phosphates and the inhibitory effects of polyvalent cations on the dephosphorylation of inositol phosphates mean that chelation can significantly affect the physical, chemical and biological processes controlling the solubilization and bioavailability of inositol phosphates. Chelating agents or ligands are complex organic anions that have multiple functional groups sharing pairs of electrons with a centrally located cation. The cations that are commonly associated with inositol phosphates are transitional metal ions, such as iron(III), zinc and copper(II). It is not within the scope of this chapter to extensively discuss coordination chemistry, but the salient concept essential to the understanding of the extractability and exchangeability of insoluble inositol phosphates in manure and soils is the relative strength of metal chelates. The formation and stability of chelating agents and cation complexes have been expressed as an equilibrium constant or the ratio of activities of the cation–ligand complex and the dissociated cation and ligand (Anderegg, 1971; Martell and Smith, 1974; Lindsay, 1979). The general

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case of ionic equilibria between a cation and a chelating ligand can be expressed as

Ligand exchange

M + nL D MLn

Organic anions and plant root exudates have long been implicated in the acquisition of phosphorus by plants grown in phosphorus-deficient soils (Ae et al., 1990; Hinsinger, 2001; Ryan et al., 2001; Jones et al., 2003). Excreted organic anions comprise a variety of products of the citric acid cycle, bearing one or more –COO− functional groups that exchange with phosphate anions and chelate the counterions. For example, piscidic acid (p-hydroxybenzyl tartaric acid) and its p-methoxybenzyl derivative are released in the rhizosphere of pigeon pea [Cajanus cajan (L.) Millsp.], allowing the legume to utilize iron-bound phosphate. In soil, commonly identified low-molecular weight organic anions include formate, acetate, propionate, oxalate and citrate, which are formed during microbial metabolism and decomposition of plant residues (Rovira and McDougall, 1967). Soil organic matter contributes aliphatic acids, phenols, phenolic acids, fulvic and humic substances (Ritchie et al., 1982; Freche et al., 1992; Haynes and Mokolobate, 2001). These humic matter ligands are large and have multiple functional groups, binding cations and releasing soluble phosphorus to the soil solution. Animal manure and wastewater also contain large amounts of organic matter that likely generate complex ligands (Ma et al., 2001). Mechanisms of phosphate and inositol phosphate solubilization in liquid manures may include ligand exchange, complexation of counterions and solubilization (Earl et al., 1979; Jones and Darrah, 1994; Kirk et al., 1999). Organic ligands have been shown to inhibit the precipitation of di-, tri- and octacalcium phosphates or hydroxyapatite, forming phospho–citrate complexes that inhibit the precipitation reaction in domestic wastewaters (Sharma et al., 1992; House, 1999; van der Houwen and Valsami-Jones, 2001). The exchangeability of ligands thus forms the basis for the development of an approach to enhancing the efficiency of enzymatic methods to determine bioavailability and solubility of phosphate and inositol phosphates in manures (Dao, 2004b).

(11.1)

where n = 1, 2, 3, . . ., i. A formation constant for the MLn species is defined as K ML n =

[ML n ] [M] [L]n

In systems of multidentate ligands such as myoinositol hexakisphosphate and polycarboxylate ligands (e.g. 1,2-cyclohexanediamine tetraacetate diaminocyclohexane tetraacetate (CDTA) and ethylenediaminetetraacetate (EDTA)), the formation of hydrogen complexes also accompanies the cation chelation process in which one or more of the donor atoms of the ligand are linked to a proton. The multidentate ligand forms protonated species: H + L D HL D H 2 K D gH p L As such, the protonation equilibria must also be defined as pH + L D H p L

(11.2)

yielding stepwise protonation constants: K p=

[H p L] [H] [H p - 1 L]

Combining Equation (11.1) and Equation (11.2), the overall stability constant of the cation chelate is expressed as K ML n =

[MH p L n ] [M] [H]p [L]n

(11.3)

A comparison of the likelihood of a cation–ligand complex to remain intact can be made between pairs of cations and ligands and to determine whether an exchange between two specific ligands would occur. An example of calculations of charge concentrations and exchange between myo-inositol hexakisphosphate and polycarboxylate ligands can be found elsewhere (Dao, 2004b). Phase diagrams of equilibrium relationships between cations and ligands as a function of pH and oxidation–reduction potential can provide useful guidelines for developing extracting conditions that favour the solubilization and exchange of phosphate and organic inositol phosphate anions with comparable or stronger ligands. The considerable body of literature on the role of organic acids in the exchange with sorbed phosphate has been reviewed (Mu et al., 1995; Jones et al., 2003).

Effects of ligand/phosphorus ratios on phytase activity Owing to the ability of one ligand to participate in exchange reactions with another ligand, a detailed study of the effects of polydentate ligands

Ligand Effects in Animal Manures

on the dephosphorylation of myo-inositol hexakisphosphate was conducted in the presence of calcium, aluminium and iron at levels that inhibited the enzymatic dephosphorylation reaction. Polydentate ligands of different sizes and charge characteristics included CDTA, diethylene triaminepentaacetate (DTPA), EDTA, oxalic acid (ethanedioic acid) and phthalic acid (1,2-benzenedicarboxylic acid). They were assessed for their ability to decouple complexed myo-inositol hexakisphosphate, thus allowing enzyme binding and dephosphorylation to occur. Effects of ligand/myo-inositol hexakisphosphate molar ratio It was first established that EDTA and the other ligands did not appear to interfere with the hydrolytic activity of phytases at molar concentrations up to 6.1 and 22.6 mM for EDTA and phthalate, respectively, when no polyvalent cation was present (Figs 11.2a and 11.3a). However, there were contradictory reports of organic ligands inhibiting the activity of phytases and broadspectrum phosphatases. Polyvalent anions such as arsenate, molybdate, L-tartrate and phosphate have been found to inhibit the activity of acid phosphatases, while EDTA, oxalate and citrate suppressed yeast (Saccharomyces cerevisiae) phytases but did not affect the activity of wheat (T. aestivum L.) bran phytases (Nayini and Markasis, 1984, 1986). However, the presence of proteins, ligands and inositol phosphates can interfere with colorimetric measurements of phosphate and underestimate the rate of reaction and extent of dephosphorylation of inositol phosphates (Shand and Smith, 1997; He et al., 1998; Dao, 2003). Therefore, the apparent contradictory evidence may have been an artefact of the analytical methodology at ligand concentrations above critical levels. Adding ligands to a mixture of myo-inositol hexakisphosphate and aluminium reversed the inhibitory effect of aluminium ions on the dephosphorylation of the inositol phosphate. When aluminium concentrations exceeded 0.75 mM, the extent of this phenomenon was in the following order: phthalate = oxalate < DPTA < EDTA = CDTA. The ligands CDTA and EDTA were able to completely reverse the inhibition of inositol phosphate hydrolysis at all aluminium concentrations up to 1.5 mM. At pH

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4.5, CDTA and EDTA were present primarily as divalent H2CDTA2− and trivalent HEDTA3−, respectively. They were both tetradentate and were apparently comparable in reactivity and ability to decouple counterion and myo-inositol hexakisphosphate for phytases to hydrolyse the latter compound. The ligands were not as effective in reversing the inhibitory effects of polyvalent cations at pH 6. Overall, EDTA was much more effective than CDTA, particularly at aluminium concentrations >0.75 mM (Fig. 11.3). In fact, EDTA was the only ligand that reversed the inhibition of the dephosphorylation reaction when the aluminium concentration was 1.5 mM. These results indicated that charge concentration was a major factor in countering the inhibition of dephosphorylation by the cation (see later). Charge concentrations for EDTA4− and the chelating ability exceeded those of CDTA, which theoretically existed as H2CDTA2− and HCDTA3−. Furthermore, aluminium existed as both Al(OH)2+ and Al(OH)3 (aq) at the higher pH. Amorphous metal hydroxides sorbed and shielded myo-inositol hexakisphosphate from dephosphorylation. Meanwhile, oxalate, commonly used to chelate iron and aluminium, was only able to partially reverse the inhibition of dephosphorylation at the 0.25 mM aluminium treatment and oxalate/myo-inositol hexakisphosphate molar ratios ≥3:1 (Fig. 11.3). Oxalate was completely ineffective at higher aluminium concentrations and was even less effective at pH 6, where it should exist as the fully dissociated conjugate base. Effects of ligand/myo-inositol hexakisphosphate charge concentration ratio The polycarboxylate ligands under study possessed two (oxalate and phthalate), four (CDTA and EDTA) or five (DTPA) carboxylate functional groups. Increasing charge concentrations reversed the inhibitory effect of polyvalent cations (Figs 11.2 and 11.3). The susceptibility to hydrolysis increased linearly with ligand/myoinositol hexakisphosphate charge concentration ratios between 1 and 4. The tetra- and pentadentate EDTA, CDTA and DTPA were most able to dissociate aluminium and myo-inositol hexakisphosphate, and increased the dephosphorylation of the dissociated anion.

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Fig. 11.2. Effect of polycarboxylate ligands, ligand/myo-inositol hexakisphosphate mole ratios, and solution pH on the dephosphorylation of inositol hexakisphosphate by Aspergillus ficuum phytase at four levels of aluminium counterion a=pH 4.5; b=pH 6.0. (A = 0; B = 0.25; C = 0.75; and D = 1.5 mM). (Reprinted from Dao, 2004b.)

Excess charge was needed to decouple and mobilize the complexed myo-inositol hexakisphosphate, and to attain the ligand/myo-inositol hexakisphosphate charge concentration ratios between 1 and 4, equivalent molar concentration ratios had to be between 1.5- and 12-fold that of myo-inositol

hexakisphosphate, except in the case of phthalate. However, there must be an upper limit to increasing charge concentration needed to overcome the inhibitory effect of polyvalent counterions, because molar concentrations should eventually reach levels that would precipitate the enzyme.

Ligand Effects in Animal Manures

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Fig. 11.3. Effect of organic ligands, ligand/myo-inositol hexakisphosphate charge ratios and equivalent molar ratios on the dephosphorylation of myo-inositol hexakisphosphate by Aspergillus ficuum phytase at four levels of aluminium counterion (A = 0; B = 0.25; C = 0.75; and D = 1.5 mM), at a solution pH of 4.5. (Reprinted from Dao, 2004b.)

Efficacy of polycarboxylate ligands in dairy manure suspensions An exchange between ligands and myo-inositol hexakisphosphate anions also took place when both compounds were added together to dairy manure suspensions, which enhanced the enzymatic dephosphorylation of the added organic phos-

phate. Increases in the concentration of aluminium and iron in the manure suspension predictably reduced dephosphorylation of myo-inositol hexakisphosphate. The ligands EDTA, CDTA and, to some extent, DTPA reversed the inhibitory effect of aluminium and iron(III) ions at cation/myoinositol hexakisphosphate and ligand/myo-inositol

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hexakisphosphate molar ratios ≤3:1 and an iron/myo-inositol hexakisphosphate molar ratio of 6:1 [1.5 mM iron(III)]. Meanwhile, phthalate and oxalate had even less or no effect on removing the inhibitory effect of aluminium or iron in manure suspensions compared to simple buffered solutions. The dephosphorylation of added or native inositol phosphates was indistinguishable. In manure suspensions amended with only exogenous myo-inositol hexakisphosphate, in the absence of added polyvalent counterions, CDTA and EDTA addition increased the release of phosphate over and above the added myo-inositol hexakisphosphate. This suggested that these ligands increased the solubilization and dephosphorylation of complexed organic phosphorus native to the manure and not merely the dissolution of inorganic phosphates; that is, there was no detectable effect of the ligands alone in unamended manure beyond that released as EDTAexchangeable phosphorus. The results imply that the addition of polycarboxylate organic ligands to the assay mixture allows a more complete measurement of soluble and complexed inositol phosphate in animal manures. Differentiating pools of phosphate and inositol phosphates in animal manure Phosphorus exists in a number of chemical forms that depend upon feed composition, extent of mineral supplementation in the feed, feed intake and absorption efficiency as well as the external physical conditions upon excretion. A mild ligand-based fractionation assay has been developed to differentiate bioavailable phosphate and phosphate monoesters, including inositol phosphates in manure (Dao, 2003; Dao et al., 2006) and soils (Dao, 2004a; Dao et al., 2005), into pools that reflect their potential solubilization. The task is critical to the accurate assessment of the fate of manure phosphorus in the environment, including the contribution of manure-derived inositol phosphates to eutrophication. Fractions of manure phosphorus that are measured in the PHP assay include: (i) waterextractable phosphate; (ii) an EDTA-exchangeable phosphate pool that was not previously extracted by water alone; (iii) a water-extractable organic phosphorus pool that is hydrolysable by phytase; (iv) an EDTA-exchangeable organic phosphorus pool that is hydrolysable by phytase;

and (v) a residual pool that is not extractable by water or ligands. Water-extractable phosphorus fraction. In animal manures, water-extractable phosphorus includes mainly soluble phosphate and small quantities of dissolved organic phosphorus species such as phosphate monoesters and nucleotides (Gerritse and Eksteen, 1978; Dou et al., 2000; He and Honeycutt, 2001; Turner and Leytem, 2004). Enzymatic assays for organic phosphoruscontaining compounds rely on the detection of released phosphate; hence the determination of water-extractable phosphate is essential to the accuracy of the methods and the effectiveness of enzymatic methods as a quantitative measurement tool for organic phosphorus. Although water-extractable phosphorus would appear to be a simple measurement, much variability and analytical artefacts exist in current methods, resulting in contradictory observations. For example, extraction periods (1–24 h or longer) and solution/solid ratios (10:1 to 200:1) vary widely. Changes in solution chemistry and abiotic factors, such as temperature, pH, oxidation– reduction potential or changes in solution/solid ratio, can alter the water-extractable phosphorus concentration based on the impact on the solubility of mineral phosphate species in manures. Inadequate knowledge of the wastewater chemistry, coordination chemistry and their effects on phosphate dissolution in complex manure mixtures may explain in part the observed variability in analytical protocols and in water-extractable phosphorus results. Another common error in the determination of water-extractable phosphorus is to overestimate the dissolved phosphate pool by not recognizing the ever-present biological processes. Concurrent phosphate-generating processes such as hydrolysis of organic phosphorus forms by manure phytases or phosphatases increases waterextractable phosphate concentrations during extraction. Therefore, water-extractable phosphate concentrations determined using longer equilibration periods would also include some phosphate released by enzymatic hydrolysis of organic phosphorus (Dao et al., 2006). A comparison of time series measurements of the liquid phase of undiluted samples and batch-diluted samples were made to illustrate the pitfalls of these laboratory estimates. These observations are reminiscent of the misinterpretation of sorp-

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tion and transformation processes affecting labile reactive organic chemicals in microbially active systems (Dao et al., 1982; Scott et al., 1982; Dao and Lavy, 1987). Common conditions that can contribute to high apparent water-extractable phosphorus include heat-drying, freeze-drying of liquid manure or extended equilibration periods that promote concurrent production of a common end product by multiple phosphate-generating pathways, including artificially enhanced biological degradation during measurement. Using a simple set of standardized protocols to minimize the latter artefact (a solid/water ratio of 1:100, w/v; 1 h equilibration), water-extractable phosphorus averaged about 15.9% ± 14.8% of manure total phosphorus, with a median value of 9.9% in a case study of 107 manure samples collected from dairy farms located across five states of the north-east USA (Lugo-Ospina et al., 2005). Water-extractable phosphorus in freshly collected manure also averaged about 37 mmol P/kg or 12.9% of the total phosphorus (Dao and Daniel, 2002) and both sets of results suggested that most of the phosphorus in dairy manure was associated with the particulate phase. LIGAND-EXCHANGEABLE INORGANIC PHOSPHATE. Tetra- or pentadentate ligands can be used to induce an exchange between the added ligand and inorganic and organic phosphates and to determine sorbed and precipitated inorganic phosphates that were not previously extracted by water alone. An additional fraction, ranging between 6.3% and 22.9% of the total phosphorus was extracted from 107 samples of dairy manure (Dao et al., 2006). The ligand forms coordination complexes with the

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released cations, preventing re-precipitation of mineral phosphates and maintaining the released phosphorus in the solution phase. The immediate measurements of phosphate concentrations in the EDTA extract yielded mainly a measure of complexed inorganic phosphates in the manures, because no exogenous phytases were used and hydrolytic enzyme kinetics were found to be slow under the experimental conditions. The sum of calcium and magnesium ions in the EDTA-extracting solution was correlated to the EDTA-exchangeable phosphorus concentration (Table 11.1). This was not unexpected, because high levels of mineral calcium are added to the diet of lactating dairy cattle (i.e. 0.37%). PHYTASE-HYDROLYSABLE PHOSPHORUS. Adding phytases to manure suspensions resulted in further increases in phosphate that were not previously extracted by water alone. The net PHP content of all samples of the manure set averaged 32.2% ± 15.6% of manure total phosphorus (as determined using a non-specific phytase). These results suggest the presence of organic phosphorus substrates that include myo-inositol hexakisphosphate and other inositol phosphate monoesters in dairy manure collected from farms across five northeastern states of the USA. Although spilled feeds and bedding materials mixed with the manures could contribute myo-inositol hexakisphosphate to the mixtures, previous work has shown that faeces and reconstituted dairy manure, prepared from freshly collected faeces and urine, also had a distinct PHP fraction (Dao, 2004b). EDTA-EXCHANGEABLE PHYTASE-HYDROLYSABLE PHOSPHORUS. Using both ligands and enzymes in

Table 11.1. Binary relationships between extractable calcium, magnesium and water-extractable and bioavailable phosphorus fractions in 107 manure suspensions collected from dairy farms across five states of the north-east USA. (Adapted from Dao et al., 2006.) Y Variable

X Variable (mol/kg)

Phosphorus fraction (mol/kg)

EDTA-extractable calcium

EDTA-extractable calcium and magnesium

Water-extractable phosphorus

y = 1.70 x + 0.033 r 2 = 0.396 y = 0.684x − 0.222x 2 − 0.007 r 2 = 0.788 y = 0.194x − 0.194x 2 + 0.037 r 2 = 0.432

y = 1.27x + 0.020 r 2 = 0.460 y = 0.488x − 0.112x 2 − 0.035 r 2 = 0.799 y = 0.088x − 0.011x 2 + 0.047 r 2 = 0.380

EDTAa-extractable phosphorus Phytase-hydrolysable phosphorus

a

EDTA = ethylenediaminetetraacetate.

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the extracting solution yielded the greatest concentration of phosphorus among any of the previous assays, because the inhibitory effect of polyvalent cations that shielded complexed myoinositol hexakisphosphate from hydrolysis was removed by the ligands. It must be recognized that the discussion is focused on EDTA, but there is no question that other polydentate ligands, e.g. CDTA, can be equally or more efficient in the ligand-exchange process. However, the theoretical basis for improvement in the efficiency of enzymatic methods for characterizing phosphorus pools in animal manure and soils remains the same. In addition, using analytical methods appropriate for the chemical species in each fraction is critical to the proper classification and mass balance of phosphorus forms in the manure sample (Dao, 2003; Dao et al., 2006). It is important to note that commercial phytase preparations used in the studies described earlier are also active against a number of phosphate monoester and diester substrates, in addition to myo-inositol hexakisphosphate (McKelvie et al., 1995; Shand and Smith, 1997; Hayes et al., 2000; Turner et al., 2002). Use of a more specific phytase preparation might confirm the identity of the derivatives of feed inositol phosphates in animal excreta. However, this would not improve the knowledge of the potential biological availability of organic phosphorus present in manure. From the perspective of agricultural and environmental sustainability, the critical issue is that animal manure contains a great deal of bioavailable organic phosphorus (i.e. compounds that can be hydrolysed by phosphatases). This means that the knowledge of all potentially bioavailable phosphorus forms better reflects the magnitude of the threat of manure phosphorus to water quality following transport in runoff to aquatic environments. Temporal changes in biologically available phosphorus The ligand-based PHP assay defined earlier provides a relative scale of chemical and biological stability of phosphorus in manures. Phosphorus pools with greater stability, whether the substrates are inorganic phosphates, inositol phosphates or other organic phosphates, may become biologically available over longer time scales in

soil. This was observed in a case study of the potential for dissolution and solubilization of immobilized phosphorus in soils treated with additives to reduce soluble phosphorus in soils (Dao et al., 2005). A freshly prepared iron hydroxide additive reduced water-extractable phosphorus in soil by 90% over a period of 16 weeks. In addition, a plant-available phosphate fraction (Mehlich-3 extractant) was also reduced in iron-treated soils, and both fractions remained unchanged up to 16 weeks. The ligand-based PHP assay, on the other hand, revealed a completely different picture of the internal changes in phosphorus pools and showed that the effect of the iron additive was transitory (Table 11.2). The inorganic EDTAextractable phosphorus fraction and the PHP pool were being remobilized, reaching initial soil levels to nullify the phosphorus-immobilizing action of the iron additives by about the fourth week following soil treatment. It also appeared that the mobilization of the PHP pool occurred as a sequential multistage process. The processes of solubilization and desorption of bioavailable phosphorus in unamended and Fe(OH)3-amended soils were best described by single or sequential double exponential kinetic equations. This behaviour would be consistent with the fact that these substances can range from meta-stable non-ordered to semicrystalline physical states (Lindsay, 1979; Zhang et al., 1992; Arai and Sparks, 2002). After 16 weeks, the total pool of potentially bioavailable phosphorus had returned to, or exceeded, the initial concentration in untreated soils, reaching an average 28% ± 1.0% and 56% ± 1.7% of total phosphorus content in the Thurmont and Burch soils, respectively. In comparison, Mehlich-3 phosphorus concentrations represented 16% ± 0.4% and 20% ± 0.3% of soil total phosphorus. Thus, the PHP assay can detect changes in the susceptibility of soil phosphorus inorganic and organic pools to transformations that traditional soil test methods do not account for.

Conclusions In areas of intensive animal production, agricultural soils have become enriched with phosphorus because of years of continuous manure applications. Organic phosphorus is an important

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Table 11.2. Kinetic models describing temporal changes in water-extractable and bioavailable phosphorus fractions in unamended soil and soils amended with iron hydroxide. (Adapted from Dao et al., 2005.) Soils Phosphorus fraction

Amendment

Thurmont gravelly loam

Burch sandy loam

Water-extractable phosphorus

Unamended

y = a + b(exp(−cx)), z where a = 9.1, b = 12.4, c = 0.989 Not fitted y = a + b(1 − exp(−cx)), where a = 109.6, b = 12.5, c = 0.0792

y = a + b(exp(−cx)), where a = 9.8, b = 15.3, c = 0.434 Not fitted y = a + b(1 + (c exp(−dx) − d exp(−cx))/(d − c)), where a = 63.9, b = 34.1, c = 0.1931, d = 0.1931 y = a + b(1 − exp(−cx)), where a = 328.6, b = 187.1, c = 0.068 y = a + b(1 + (c exp(−dx) − d exp(−cx))/(d − c)), where a = 172.9, b = 49.7, c = 0.3862, d = 0.3862 y = a(1 − exp(−bx)), where a = 165.4, b = 0.250

EDTAa-exchangeable phosphorus

+ Fe(OH)3 Unamended

+ Fe(OH)3

Phytase-hydrolysable phosphorus

Unamended

+ Fe(OH)3

y = a + b(1 − exp(−cx)), where a = −38.9, b = 117.5, c = 0.391 y = a + b(1 + (c exp(−dx) − d exp(−cx))/(d − c)), where a = −4293.8, b = 4658.6, c = 2.785, d = 2.7823 y = a + b(1 − exp(−cx)), where a = −378.4, b = 620.5, c = 0.604

a

EDTA = ethylenediaminetetraacetate.

aspect of this problem, because it represents a considerable proportion of the phosphorus in plant residues and animal manures and presents a threat to aquatic systems following its transfer in runoff from soils to water bodies. Speciation of inositol phosphates in environmental samples is poorly understood, which hampers the understanding of the environmental behaviour and transformations of inositol phosphates. It is therefore difficult to develop comprehensive strategies for managing excess nutrients in animal agricultural production systems. This chapter reviewed factors affecting the solubilization and dephosphorylation of dissolved and insoluble complexes of myo-inositol hexakisphosphate in animal manures. An in situ ligand-based enzymatic method was described, which provides insight into the biological stability of inositol phosphates in manures. Selected polydentate ligands and fungal phytases can differentiate various phosphorus pools that contribute to the solution-phase phosphate con-

centration in animal manures and manureamended soils. The biological environment, including the activity of extracellular phytases, is hard to control and varies widely across soils, agroecosystems and climatic regions. Yet it plays a key role in regulating the fate of inositol phosphates in the environment. The inclusion of biological and biochemical mechanisms in the methodology for assessing manure-derived inositol phosphates may therefore reflect more accurately their availability to microorganisms and plants. Moreover, these mechanisms can reveal the underlying potential for their dephosphorylation and release of phosphate in the long term. Further studies on the role of manure chemistry and ligands in controlling inositol phosphate solubilization and mobility will improve our understanding of the linkage between watersoluble, exchangeable and hydrolysable forms, and their mobility in soils and the wider environment.

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Wise, A. and Gilburt, D.J. (1981) Binding of cadmium and lead to the calcium-phytate complex in vitro. Toxicological Letters 9, 45–50. Zhang, J., Ebrahimpour, A. and Nancolas, G.H. (1992) Dual constant composition studies of phase transformation of dicalcium phosphate dihydrate into octacalcium phosphate. Journal of Colloid Interface Science 152, 132–140. Zhang, H., Dao, T.H., Basta, N.T., Dayton, E.A. and Daniel, T.C. (2002) Remediation techniques for manure nutrient loaded soils. National Center for Manure and Animal Waste Management. White Papers on Animal Agriculture and the Environment CDROM No WP-CD-1, Chapter 13. Midwest Plan Service, Ames, Iowa. Zyla, K., Wikiera, A., Koreleski, J., Swiatkiewicz, S., Piironen, J. and Ledoux, D.R. (2000) Comparison of the efficacies of a novel Aspergillus niger mycelium with separate and combined effectiveness of phytase, acid phosphatase, and pectinase in dephosphorylation of wheat-based feeds fed to growing broilers. Poultry Science 79, 1434–1443.

12

Inositol Phosphates in Soil: Amounts, Forms and Significance of the Phosphorylated Inositol Stereoisomers Benjamin L. Turner Smithsonian Tropical Research Institute, Apartado 0843-03092, Balboa, Ancón, Republic of Panama

Inositol phosphates are abundant in soil, yet their origin, dynamics and ecological function remain largely unknown. Given the importance of phosphorus in agriculture and the environment, it is remarkable that so little is known about one of the most prevalent forms of soil organic phosphorus. In particular, several inositol phosphates that are common in soil occur nowhere else in nature. Inositol phosphates were first reported in soil more than 60 years ago (Dyer et al., 1940; Yoshida, 1940) and their quantitative importance soon became apparent. Numerous reports in the following decades, including the pioneering work of Dennis Cosgrove and George Anderson, documented the amounts and forms of inositol phosphates in soil. However, there has been little additional research since Michael L’Annunziata’s studies of inositol phosphate stereochemistry in the mid-1970s (see L’Annunziata, Chapter 4, this volume). This has been due in part to the widespread adoption of solution 31P nuclear magnetic resonance (NMR) spectroscopy as the method of choice for the analysis of soil organic phosphorus, which conventionally provides data on phosphate monoesters as a broad functional group rather than as individual compounds such as the inositol phosphates. However, recent advances in methodology mean that detailed information on soil inositol phosphates can now be obtained using standard NMR procedures (Turner et al., 2003b; Turner and Richardson, 2004), which should reinvigorate research on the inositol phosphates in soil. 186

Other chapters in this volume deal specifically with the reactions, mobility, hydrolysis and bioavailability of inositol phosphates in the environment. This chapter addresses specifically the amounts, forms and functions of inositol phosphates in soil. Emphasis is placed on the phosphorylated stereoisomers, some of which have never been detected in biological tissue. The chapter builds on a recent review of inositol phosphates in the environment (Turner et al., 2002) and the reader is also referred to a series of other reviews of soil organic phosphorus that include information on the inositol phosphates (Anderson, 1967; Cosgrove, 1967; Halstead and McKercher, 1975; Dalal, 1977; Anderson, 1980; Cosgrove, 1980; Harrison, 1987; Stewart and Tiessen, 1987; Magid et al., 1996; Condron et al., 2005).

Amounts of Inositol Phosphates in Soil There is a considerable body of data on inositol phosphates in soil, some of which is summarized in Table 12.1. From this it seems reasonable to conclude that the inositol phosphates are quantitatively important in soil, although it is also clear that their concentrations and contribution to the soil organic phosphorus vary widely. This is illustrated in Fig. 12.1, which shows solution 31P NMR spectra of the phosphorus extracted from

©CAB International 2007. Inositol Phosphates: Linking Agriculture and the Environment (eds B.L. Turner, A.E. Richardson and E.J. Mullaney)

Inositol Phosphates in Soil

Phospholipid degradation products

(a)

187

RNA mononucleotides Pyrophosphate

5

4

DNA

5

(b)

−5

0

myo-Inositol hexakisphosphate

Phosphate

scyllo-Inositol hexakisphosphate

7

5

6

0

5

4

3

−5

Chemical shift (ppm) Fig. 12.1. The organic phosphorus composition of two soils from the USA with contrasting proportions of inositol phosphates determined by extraction in sodium hydroxide and ethylenediaminetetraacetate (EDTA) and solution 31P nuclear magnetic resonance (NMR) spectroscopy. (a) The spectrum of a wetland soil from a nutrient-enriched site in the Florida Everglades that contained no detectable inositol phosphates. (From Turner and Newman, 2005.) The soil was a Histosol containing 44% carbon and 0.16% phosphorus. (b) The spectrum of an arable soil from Sussex County, Delaware, in which all detectable organic phosphorus in the extract was inositol hexakisphosphate. (From P. Murphy and B. Turner, 2003, unpublished data.) The soil was an acidic sandy loam containing 0.7% carbon and 0.09% phosphorus. The zoomed inset spectra show the phosphate monoester regions in detail. The main spectrum in (a) is plotted with 8 Hz line broadening, while all other spectra are plotted with 1 Hz line broadening to show fine resolution.

two US soils. One soil contains virtually all its organic phosphorus in the form of inositol phosphates, while the other contains none. It should not therefore be assumed that inositol phosphates are abundant in all soils. The majority of the inositol phosphates in soil are hexakisphosphates (i.e. with six phosphate

groups around the inositol ring). This is probably due to a combination of factors, but principally that most inputs to soil are as myo-inositol hexakisphosphate from plants. The pentakisphosphates are also abundant in some soils, altough lowerorder esters (monophosphates to tetrakisphosphates) are relatively rare (Omotoso and Wild,

188

Table 12.1. Concentrations of inositol hexakisphosphate in surface soil from various parts of the world.

Location

Anderson (1964)

17

Scotland

Caldwell and Black (1958c)

49

USA

Islam and Mandal (1977)

20

McKercher and Anderson (1968b) Thomas and Lynch (1960)

Reference

Soil type and land use

Soil properties

Range of textural properties, mainly loams; land use not specified Range of virgin and cultivated soils

Carbon 1.0–8.0% pH 4.7–7.5 Organic Pb 220– 920 mg P/kg pH 4.9–7.8 Organic Pb 40– 446 mg P/kg

Bangladesh

Range of Ultisols, Entisols and Inceptisols; land use not specified

18

Canada

Range of soils; land use not specified

9

Canada

Virgin loams

Organic matter 0.7–3.6% pH 4.7–8.1 Organic Pb 63– 261 mg P/kg Nitrogen 0.08–0.59%e Organic Pb 100–475 mg P/kge Organic matter 2.4–13.6% pH 5.5–7.1 Organic Pb 148–710 mg P/kg

Inositol hexakisphosphate (mg P/kg)

Proportion of the total organic P (%)a

Hot 3 M NaOH, ion-exchange chromatography

56–460

24–58 (40)

Concentrated HCl, 0.5 M NaOH, ion-exchange chromatography Hot 3 M NaOH, ion-exchange chromatography

2–62c

2–31 (12)

19–130d

21–58 (36)

Hot 3 M NaOH, ion-exchange chromatography

20–71d

11–23 (17)

Concentrated HCl, 0.5 M NaOH, ion-exchange chromatography

6–72c

2–10 (5)

Analytical procedure

B.L.Turner

Number of soils

Turner (2006)

13

Madagascar

Humid tropical Oxisols under rice

Turner et al. (2003b)

29

England and Wales

Williams and Anderson (1968)

47

Australia

Temperate lowland permanent pasture with high clay (22–68%) Range of soils and land use, including cultivated, uncultivated and pasture

NaOH–EDTA ND–33c extraction, solution 31P NMR spectroscopy

ND–26

NaOH–EDTA 26–189c 31 extraction, solution P NMR spectroscopy

11–35 (22)

1–356d

<1–38 (16)

Hot 3 M NaOH, ion-exchange chromatography

Inositol Phosphates in Soil

ND = not detected; NMR = nuclear magnetic resonance. a Values in parentheses are the mean of all soils. b Determined by the extraction method of Mehta et al. (1954). c Values are for myo-inositol hexakisphosphate only. d Includes inositol pentakisphosphates. e Values estimated from figure. f Determined by NaOH–EDTA extraction and solution 31P NMR spectroscopy. g Determined by the extraction method of Saunders and Williams (1955).

Carbon 1.1–15.3% pH 4.6–5.8 Organic Pf 22–393 mg P/kg Carbon 2.9–8.0% pH 4.4–6.8 Organic Pf 208–895 mg P/kg Nitrogen 0.04–0.76% pH 5.0–9.0 Organic Pg 6–1773 mg P/kg

189

190

B.L.Turner

1970a; Anderson, 1980). In fact, it remains unclear whether the lower esters occur naturally in soil or are an artefact of the strong chemical solutions necessary to extract inositol phosphates from soil (Cosgrove, 1980). The early literature contains many reports of inositol phosphates in soil, but these must be assessed with care. Most studies prior to the 1980s involved strong alkali extraction followed by column chromatography, which may have inaccurately determined the soil inositol phosphate content (Turner et al., 2002). Problems included the incomplete recovery of inositol phosphate from anion-exchange columns (Anderson, 1964; Martin, 1970) and the inclusion of organic phosphorus compounds other than inositol phosphates in the separated fractions (Irving and Cosgrove, 1981). These problems are exemplified by discrepancies observed in studies that compared the various methods. Dormaar (1967) measured greater inositol phosphate concentrations in Canadian chernozems using the method of Caldwell and Black (1958a) compared with a procedure based on the methods of Anderson (1956) and Cosgrove (1963). In direct contrast, McKercher and Anderson (1968b) recorded less inositol phosphate in Canadian soils by the Caldwell and Black (1958a) method compared with the method of Anderson (1964). The authors suggested that this might have been caused by incomplete precipitation of iron-phytate when using the wide iron/inositol phosphate ratio of the Caldwell and Black (1958a) procedure. However, Anderson (1964) had previously reported little difference between his method and that of Caldwell and Black (1958a) for a range of British soils. Recent studies employed a single-step alkaline extraction procedure with detection by solution 31P NMR spectroscopy (Turner et al., 2003b, 2005b; Turner and Richardson, 2004). Soil organic phosphorus recovery in a solution containing sodium hydroxide and ethylenediamine tetraacetate (EDTA) is similar to the conventional methods used in the older literature (Bowman and Moir, 1993; Turner et al., 2005a), so presumably results from recent and earlier studies are broadly comparable. Using solution 31P NMR spectroscopy, it is currently possible to quantify myoinositol hexakisphosphate by its four characteristic signals that occur in a 1:2:2:1 ratio (Turner et al., 2003b), while scyllo-inositol hexakisphosphate is

identified by its single strong signal at the upfield end of the phosphate monoester region (all six phosphate groups are stereochemically identical) (Turner and Richardson, 2004). The signals can be determined directly in extracts by spectral deconvolution or following hypobromite oxidation (Fig. 12.2). The latter procedure destroys all phosphate monoesters except the inositol phosphates (Wrenshall and Dyer, 1941) and conveniently improves the spectral resolution if EDTA is included in the NMR tube (Fig. 12.2; Turner and Richardson, 2004). Given that detection by NMR spectroscopy precludes the problems associated with column chromatography, NaOH–EDTA extraction and solution 31P NMR spectroscopy provide a convenient and accurate alternative to the conventional procedures for the determination of inositol phosphates in soil. Further improvement is possible by using two-dimensional NMR spectroscopy (see Murthy, Chapter 2, this volume), although this has not yet been applied to the complex matrices of soil extracts.

Factors Controlling the Amounts of Inositol Phosphates in Soil Despite the large amount of published information on inositol phosphates in soil, there is still no clear understanding of the factors controlling their abundance. Indeed, the proportion of the soil organic phosphorus as inositol phosphates can vary appreciably, even in nearby soils formed under similar environmental conditions and derived from the same parent material (Williams and Anderson, 1968; Turner et al., 2003b). In a meta-analysis of literature information, Harrison (1987) reported that ~90% of the variation in the concentrations inositol phosphates in soils was explained by organic phosphorus, pH and organic carbon. This is relatively uninformative, however, given that inositol phosphates typically constitute most of the soil organic phosphorus. Studies of a large number of soils under similar climate and vegetation have indicated that inositol phosphates are not correlated with factors typically associated with organic matter content, such as organic carbon, nitrogen, clay or microbial biomass. Rather, they are correlated with factors linked specifically to phosphate stabilization, such as the phosphate sorption capacity

Inositol Phosphates in Soil

(a)

Phosphate

(b)

myo-Inositol hexakisphosphate

191

scyllo-Inositol hexakisphosphate

Unidentified inositol phosphates

(c)

7

6

5

4

Chemical shift (ppm) Fig. 12.2. Identification of scyllo- and myo-inositol hexakisphosphates in a soil extract by hypobromite oxidation and solution 31P nuclear magnetic resonance (NMR) spectroscopy. (From Turner and Richardson, 2004.) (a) The spectrum of an untreated sodium hydroxide/ethylenediaminetetraacetate (EDTA) extract of a Welsh pasture soil; (b) the same extract after hypobromite oxidation to destroy all phosphate monoesters other than higher-order inositol phosphates; (c) the same extract analysed without EDTA in the redissolved sample. Note the marked difference in resolution in the three spectra. In spectrum (b) the strong signal at 6.2 ppm is inorganic phosphate, while all other signals are inositol phosphates. The soil was a clay loam containing 4.6% carbon and 0.11% phosphorus.

and amorphous aluminium and iron (McKercher and Anderson, 1968b; Anderson et al., 1974; Turner et al., 2003b). This is unsurprising, because inositol hexakisphosphates are rapidly and strongly sorbed to soil constituents and

can form insoluble precipitates with polyvalent cations (Jackman and Black, 1951; Anderson and Arlidge, 1962). These abiotic processes account for the characteristic dynamics of inositol phosphates in the environment and are reviewed in

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B.L.Turner

detail elsewhere in this volume (see Celi and Barberis, Chapter 13). It therefore seems likely that inositol phosphates are stabilized in soil by different mechanisms to those responsible for the stabilization of organic matter. This may explain in part the discrepancies in nutrient stoichiometry observed in soil (McGill and Cole, 1981). However, at least some inositol phosphates are intimately associated with organic matter, because they have been detected in high-molecular weight humic substances isolated from soil (Moyer and Thomas, 1970; Omotoso and Wild, 1970b). In addition, the proportions of phosphate monoesters in humic acids of lowland rice soil of the Philippines were positively correlated with those of aromatic carbon and heterocyclic nitrogen, suggesting that inositol phosphates were contained within strongly humified structures (Mahieu et al., 2002). There is a decline in organic phosphorus when virgin soil is brought under cultivation (Tiessen et al., 1983), although inositol phosphates appear to be depleted at a slower rate than other organic phosphates (Williams and Anderson, 1968; Condron et al., 1990). This selective mineralization means that cultivated soils tend to contain a greater proportion of their organic phosphorus as inositol phosphates (see Fig. 12.1). Some of the differences may be explained by sampling depth, because inositol phosphates accumulate at the surface of undisturbed soil, but will be distributed throughout the plough layer in cultivated soil (Thomas and Lynch, 1960; Dormaar, 1967; McKercher and Anderson, 1968b). They can, however, accumulate in the B-horizon of podzols (Williams and Anderson, 1968). Several studies reported that temperate forest soils contained more inositol phosphates than grassland soils, both in terms of concentration and as a proportion of the soil organic phosphorus (Caldwell and Black, 1958c; McKercher and Anderson, 1968b). Those studies rarely reported the type of forest, but this is likely to be important. For example, conversion from pasture to pine forest in New Zealand caused a marked decrease in soil organic phosphorus (Condron et al., 1996), which was subsequently linked to the degradation of inositol phosphates (Chen et al., 2004). It was recently reported that inositol phosphates were absent from organic wetland soils of the Florida Everglades, USA (Turner and

Newman, 2005). As samples were analysed from several locations with a range of chemical properties, this suggests a major difference in the phosphorus cycle compared to mineral soils. Similar results were reported for constructed treatment wetlands designed to sequester pollutant phosphorus from agricultural runoff (Turner et al., 2006). This raises important concerns about the long-term stability of the organic phosphorus sequestered in such systems, because it occurs in relatively unstable phosphate diesters rather than recalcitrant inositol phosphates. Although there are several possible explanations for the absence of inositol phosphates in these wetland soils, the most likely is that anaerobicity occurs close to the sediment surface for much of the year. In acidic rice soil the hydrolysis of inositol phosphates begins soon after submergence (Islam and Ahmed, 1973), while in marine sediments hydrolysis proceeds more rapidly under anaerobic conditions than in parallel samples incubated aerobically (Suzumura and Kamatani, 1995). This may be due to the reduction of ferric iron and the release of associated inositol phosphates, which are then available for biological attack. However, anaerobic reduction of ferric iron–inositol hexakisphosphate complexes was reported to form insoluble Fe4-phytate rather than release free inositol hexakisphosphate (De Groot and Golterman, 1993). This could account for the observed decreases in inositol phosphate following submergence of rice soils, but only if, as seems unlikely, the Fe4-phytate complex remains insoluble in the strong alkali used to extract inositol phosphates. This is also discussed elsewhere in this volume (see Celi and Barbaris, Chapter 13, and McKelvie, Chapter 16). Soil pH should exert a strong control on the accumulation of inositol phosphates, because sorption and metal precipitation are greater under acidic conditions ( Jackman and Black, 1951; Anderson and Arlidge, 1962). In addition, phytases exhibit marked differences in behaviour depending on pH – they are most active under acidic conditions, but are also inactivated by sorption to a greater extent in more acidic soil (see George et al., Chapter 14, this volume). Soil organic phosphorus concentrations are often negatively correlated with soil pH (Harrison, 1987), although this is less clear for the inositol phosphates. Caldwell and Black (1958c) reported a significant negative correlation between soil pH

Inositol Phosphates in Soil

and the proportion of the soil organic phosphorus as inositol phosphates, although other studies (e.g. Williams and Anderson, 1968) found no such relationship. Inositol phosphates will readily form calcium precipitates under alkaline conditions (see Celi and Barberis, Chapter 13, this volume), but there is no clear evidence that the presence of carbonates influences inositol phosphate accumulation. Cosgrove (1966) reported the complete absence of inositol phosphates in slightly alkaline calcareous soils from Scotland, but other studies have measured considerable quantities in calcareous soils (McKercher and Anderson, 1968b; Williams and Anderson, 1968). These discrepancies may be explained by methodology, because Anderson (1964) reported a marked difference in the concentration of inositol phosphates extracted from a calcareous soil by two different procedures. There is also evidence for the relative solubility of inositol phosphates in calcareous soils. In a study of a range of mainly cultivated calcareous soils from the semiarid western USA, concentrations of phosphate monoesters determined by solution 31P NMR spectroscopy of alkaline soil extracts were relatively small and not significantly correlated to carbonate content, although they were correlated with other soil properties, including organic carbon, amorphous iron and aluminium, pH and clay (Turner et al., 2003a). However, phytase-hydrolysable phosphorus assays demonstrated that a large proportion of the organic phosphorus in bicarbonate extracts was inositol hexakisphosphate, suggesting the relative solubility and potential bioavailability of inositol phosphates in these calcareous soils. This was confirmed by a recent study that demonstrated the rapid decomposition of myo-inositol hexakisphosphate in manure within weeks of application to a calcareous arable soil (see Leytem and Maguire, Chapter 10, this volume). A key factor in the regulation of inositol phosphates in soil that has not been adequately addressed is the role of nutrient status. Inositol phosphates are conventionally considered to be relatively unavailable to organisms due to their strong interaction with soil components (see Richardson et al., Chapter 15, this volume), but many soil microbes synthesize phytase and can utilize inositol phosphates in their environment (e.g. Richardson and Hadobas, 1997; Unno et al., 2005; see Hill and Richardson, Chapter 5, this

193

volume). This raises the possibility that inositol phosphates are used only when phosphorus is scarce relative to other nutrients. In other words, phosphorus limitation may drive the biological degradation of recalcitrant inositol phosphates by favouring organisms that can access them in soil. Evidence in support of this hypothesis is limited, but a study of a large number of soils under similar vegetation and climate (temperate permanent pasture) revealed that inositol phosphate concentrations were greatest in soils with a low nitrogen/organic phosphorus ratios (Turner et al., 2003b, 2005b). In the case of myo-inositol hexakisphosphate, concentrations were also greater in soils containing more phosphate in readily soluble form (as determined by bicarbonate extraction). In agricultural soils, phosphate fertilization would be expected to decrease the biological demand for phosphorus associated with soil organic matter and lead to an accumulation of inositol phosphate. This may explain in part the abundance of inositol phosphates in cultivated soils and their persistence compared with other organic phosphates such as nucleic acids and phospholipids during cultivation (Williams and Anderson, 1968; Condron et al., 1990). Analysis of soils of different ages from the Franz Josef glacial chronosequence in New Zealand revealed a rapid increase in inositol phosphates (in terms of both concentration and proportion of the soil organic phosphorus) during the first 1000 years of soil development, after which the concentrations declined (Baker, 1977). As the early stages of ecosystem development are typically associated with nitrogen limitation (Walker and del Moral, 2003), the abundance of inositol phosphates could be linked to the plentiful availability of phosphorus. The role of nutrient status therefore deserves careful attention in future studies of inositol phosphates in soil.

Phosphorylated Inositol Stereoisomers in Soil Perhaps the most intriguing aspect of inositol phosphates in soil is the presence of stereoisomers other than myo-inositol that occur nowhere else in nature. Nine stereoisomers of inositol exist (see Shears and Turner, Chapter 1, this volume),

194

B.L.Turner

although only four have been detected in soil in the phosphorylated form (Fig. 12.3). Of these, only myo-inositol phosphates are prevalent in nature (Raboy, 2003). The remaining three stereoisomers (neo-inositol, D-chiro-inositol and scyllo-inositol), which differ from myo-inositol by the orientation of a single hydroxyl group, occur widely in soil in highly phosphorylated forms, yet are extremely rare in biological tissue (discussed below). Studies reporting quantitative estimates of the phosphorylated inositol stereoisomers in soil are summarized in Table 12.2. In agreement with the results for total inositol phosphates, the concentrations of phosphorylated stereoisomers and their contribution to the soil organic phosphorus vary widely. The dominant phosphorylated stereoisomer in soil is myo-inositol, as would be expected from its widespread abundance in plants. The remaining isomers tend to occur in the following order of abundance: scyllo-> D-chiro-> neo-, although myo- and scyllo-inositol hexakisphosphates together account for most of the soil inositol phosphate (McKercher and Anderson, 1968a; Turner and Richardson, 2004). All three stereoisomers have been detected as lower esters in small concentrations (e.g. Halstead and Anderson, 1970). It should be noted, however,

that some studies did not detect scyllo-inositol phosphates, despite the presence of other phosphorylated stereoisomers (Martin and Wicken, 1966; L’Annunziata and Fuller, 1971), while Irving and Cosgrove (1982) suggested that neoinositol phosphates are not quantitatively determined unless a hypobromite oxidation step is included in the analytical procedure. This means that neo-inositol phosphates may have been underestimated in parts of the older literature. Early studies involving chromatographic separation of inositol phosphates in soil extracts documented a ‘supposed isomer’ of myo-inositol hexakisphosphate that eluted at the end of the chromatogram (Smith and Clark, 1951; Caldwell and Black, 1958a) and was subsequently shown to be scyllo-inositol hexakisphosphate (Cosgrove, 1962). Of the few studies in which quantitative values were obtained for a large number of soils, Caldwell and Black (1958c) reported scyllo-inositol hexakisphosphate (although at the time of publication they did not identify it as such) to constitute an average of 5.8% of the organic phosphorus in US soils, while concentrations in temperate pasture soils from England and Wales averaged 9.7% of the soil organic phosphorus (Turner et al., 2005b). Concentrations of scyllo-Inositol

myo-Inositol HO

HO

HO

OH

HO

OH

OH

OH

HO HO

1 axial group

No axial groups

neo-Inositol

D-chiro-(+)-Inositol

HO

OH

OH

HO

HO OH

OH HO

HO

HO

OH

OH

OH

OH 2 axial groups

2 axial groups

Fig. 12.3. The four inositol stereoisomers that occur in phosphorylated forms in soil. Axial hydroxyls are shown in bold and hydrogen groups have been omitted. The four isomers differ only by the orientation of a single hydroxyl group (circled). Compared with myo-inositol, D-chiro- and neo-inositols have an extra axial group, whereas scyllo-inositol has none. This confers scyllo-inositol hexakisphosphate with a resistance to enzymatic attack that may account in part for its persistence in soils.

Inositol Phosphates in Soil

scyllo-inositol hexakisphosphate can exceed those of myo-inositol hexakisphosphate, although this is not common (Table 12.2). Only a handful of studies have quantified D-chiro- and neo-inositol hexakisphosphate in soil, although Irving and Cosgrove (1982) determined values in four Australian soils by gas chromatography. Ratios of D-chiro- or neo-inositol phosphates to myo-inositol phosphates are typically <0.2. Quantitative data on the phosphorylated inositol stereoisomers are fragmentary and insufficient to draw firm conclusions on factors regulating their presence and abundance in soil. Concentrations of scyllo-inositol hexakisphosphate in a series of temperate pasture soils were correlated positively with soil organic phosphorus and myo-inositol hexakisphosphate, although not with pH, organic carbon, amorphous iron and aluminium or microbial biomass (Turner et al., 2005b). However, scyllo-inositol hexakisphosphate was strongly correlated with the nitrogen/ organic phosphorus ratio, suggesting that the abundance of this compound is regulated at least in part by nutrient status. This was further supported by the results of a 10-month pot experiment with six grassland soils from New Zealand, in which the growth of ryegrass (Lolium perenne L.) decreased scyllo-inositol hexakisphosphate in three low-nutrient soils by 5–21%, but increased it in three other high nutrient soils by 11–16% (Turner et al., 2005b). This indicates that organisms that are able to access inositol phosphates may be favoured when phosphorus is relatively scarce, although further studies are clearly required to assess this in greater detail.

Origins of Phosphorylated Inositol Stereoisomers Despite their widespread occurrence in soils, the phosphorylated inositol stereoisomers other than myo have been detected rarely elsewhere in nature and their origins in soil are unknown. Inositol phosphates are conventionally considered to originate mainly from plants, but highly phosphorylated stereoisomers other than myoinositol have been detected only once in plant tissue. This suggests the importance of microbes in the synthesis of these compounds, yet they have never been detected in any soil organism.

195

Reports of the detection of inositol stereoisomers and their phosphorylated forms are summarized in Table 12.3. myo-Inositol hexakisphosphate is abundant in eukaryotes as both a cellular component and in seeds (Raboy, 2003). In contrast, scyllo-inositol hexakisphosphate has never been detected in biological tissue, although a phospholipid containing a scyllo-inositol monophosphate occurs in barley aleurone (protein stored as granules in the cells of plant seeds) (Kinnard et al., 1995; Narasimhan et al., 1997; Carstensen et al., 1999). The only known source of highly phosphorylated neo-inositol phosphates is amoebae, including the freshwater carnivorous amoeba Amoeba discoide (Laird et al., 1976) and the human intestinal parasite Entamoeba histolytica (Martin et al., 2000). In the latter organism, neoinositol occurs as both hexakisphosphate and pyrophosphate forms (i.e. with up to eight phosphate groups), although the function of these compounds remains unclear. The presence of D-chiro-inositol phosphate was reported in needles of ponderosa pine (Pinus ponderosa P. & C. Lawson) and leaves of velvet mesquite (Prosopis juliflora var. velutina (Sw.) DC. (Woot.) Sarg.) (L’Annunziata and Fuller, 1971), but it has never been detected in any other organism. An additional phosphorylated stereoisomer, muco-inositol hexakisphosphate (Fig. 12.4), was also detected in the velvet mesquite leaves and is the only report of this rare stereoisomer in any phosphorylated form in nature. However, these results were subsequently questioned on analytical grounds, because the extracts probably contained considerable polysaccharide and nitrogenous material that could have contained free inositols (Cosgrove, 1980). This is discussed in detail elsewhere in this volume (see L’Annunziata, Chapter 4). It would be appropriate to reassess similar samples using modern analytical techniques, given the potential significance of the natural occurrence of phosphorylated muco-inositol. Stereoisomeric forms of inositol hexakisphosphate were detected in aerobically digested sewage sludge in similar ratios to those detected in soils, although they constituted only around 5% of the organic phosphorus (Cosgrove, 1973). As they were not present in raw sewage, it seems likely that microbial activity was involved in their synthesis. However, only myo-inositol phosphates have been reported in manures from a wide variety of animals (including pigs, sheep, chickens,

Table 12.2. Studies reporting quantitative values for inositol hexakisphosphate stereoisomers other than myo in soils.

Reference Caldwell and Black (1958c)

Number of soils 49

Location USA

Soil type and land use

Properties

Range of virgin and cultivated soils Two samples of alpine humus and a basaltic soil

pH 4.9–7.8 Organic Pa 40– 446 mg P/kg

Not reported

Not reported

Cosgrove (1963)

3

Australia

Irving and Cosgrove (1982)

4

Australia

Martin and Wicken (1966) McKercher and Anderson (1968a)

5

New Zealand

Range of surface soils

8

Canada and Scotland

Range of arable, grassland and forest soils

Organic Pb 770–2088 mg P/kg

Carbon 6.6–12.3% Organic Pd 440–1360 mg P/kg Carbon 1.8–6.6% pH 4.9–7.8 Organic Pa 200–920 mg P/kg

Analytical procedure

Isomers detected

Concentrated HCl, 0.5 M NaOH, ion-exchange chromatography 1 M NaOH, hypobromite oxidation, ion-exchange chromatography Hot 3 M NaOH, hypobromite oxidation, gas chromatography 0.3 M KOH, ion-exchange chromatography Hot 3 M NaOH, ion-exchange chromatography

scyllo -

scyllo -

scyllo D-chiro-

neo -

Concentration (mg P/kg soil) 1–39

15–63 (scyllo)

4.2–33.1c 1.2–13.1c 1.2–10.0c

D-chiro-

scyllo D-chironeo -

9.0–87.4c

Proportion of the total organic P (%)

Ratio of isomer to myo-

1.6–21.1

0.17–1.18

2–3

0.20–0.25 (scylloto myo-+ D-chiro-)

–

0.30–0.41 0.09–0.12 0.09–0.11

4.0–42c

0.37–0.56

3.4–16.2c <5 <1

0.22–0.90 (scylloto myo- + D-chiro)

Omotoso and Wild (1970a)

7

England and Nigeria

Grassland and forest soils

Thomas and Lynch (1960)

9

Canada

Virgin loams

Turner (2006)

13

Madagascar

Turner et al. (2005b)

29

England and Wales

Carbon 1.5–13.1% pH 3.7–7.2 Organic Pa 17–175 mg P/kg

Organic matter 2.4–13.6% pH 5.5–7.1 Organic Pb 148–710 mg P/kg Humid tropical Carbon 1.1–15.3% Oxisols under pH 4.6–5.8 rice Organic Pf 22–393 mg P/kg Temperate Carbon 2.9–8.0% permanent pH 4.4–6.8 lowland pasture Organic Pf with high clay 208–895 mg P/kg (22–68%)

ND = not detected; NMR = nuclear magnetic resonance. a Determined by the extraction method of Mehta et al. (1954). b Determined by the extraction method of Saunders and Williams (1955). c Includes inositol pentakisphosphates. d Determined by the ignition method of Saunders and Williams (1955). e Values are for all esters of the stereoisomer (i.e. from monophosphates to hexakisphosphates). f Determined by NaOH–EDTA extraction and solution 31P NMR spectroscopy.

ND–5.1e 1.1–2.9e ND–0.4e

0–0.32 0.07–0.18 0–0.02

<1–12

0.45–1.83

ND–44

ND–11

0.33–0.67

11–130

4.4–14.5

0.29–0.79

1 M NaOH, hypobromite oxidation, ion-exchange chromatography Concentrated HCl, 0.5 M NaOH, ion-exchange chromatography

scylloD-chironeo -

scyllo -

2.8–83.4

NaOH–EDTA extraction, solution 31P NMR spectroscopy NaOH–EDTA extraction, solution 31P NMR spectroscopy

scyllo -

scyllo-

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Table 12.3. The occurrence of phosphorylated and free inositol stereoisomers in nature. Isomer

Axial groups Occurrence in phosphorylated form

scylloInositol

0

Common in soils as the hexakisphosphate and lower esters (Cosgrove, 1962); also detected in aerobic sewage sludge (Cosgrove, 1973); a phospholipid containing scylloinositol monophosphate occurs in barley aluerone cells (Kinnard et al., 1995)

myo-Inositol

1

D-chiro-

2

Abundant as the hexakisphosphate and lower esters in a range of organisms (Raboy, 2003) and soils (Turner et al., 2002) Abundant as the hexakisphosphate in soil, with smaller amounts of the lower esters (Cosgrove, 1969); also detected in aerobic sewage sludge (Cosgrove, 1973), pine needles and velvet mesquite leaves (L’Annunziata and Fuller, 1971) Never detected in phosphorylated form

Inositol

L-chiro-

2

Inositol

neo-Inositol

2

epi-Inositol muco-Inositol

2 3

allo-Inositol cis-Inositol

3 3

Occurs in soils as the hexakisphosphate and occasionally lower esters (Cosgrove and Tate, 1963); also detected in aerobic sewage sludge (Cosgrove, 1973); highly phosphorylated forms occur in amoebae living in human intestines (Martin et al., 2000) and fresh water (Laird et al., 1976); also present as the monophosphate in mammalian brain tissue (Sherman et al., 1971) None Detected once in velvet mesquite leaves (L’Annunziata and Fuller, 1971)

None None

Occurrence of the free inositol Widespread in plants, including chrysanthemums (Ichimura et al., 2000) and Proteaceae (Bieleski and Briggs, 2005); also detected in the ciliate Tetrahymena vorax (Kersting et al., 2003), the red algae Porphyra umbilicalis and a number of other organisms (Posternak, 1965) Widespread in biological tissue (Morré et al., 1990; Loewus and Murthy, 2000) Occurs in plants, often in methylated form as pinitol (Morré et al., 1990); also detected in trace amounts in the ciliate Tetrahymena vorax (Kersting et al., 2003) Occurs in plants such as seagrass (Drew, 1983), chrysanthemums (Ichimura et al., 2000) and Proteaceae (Bieleski and Briggs, 2005), often in methylated form Occurs in mammalian tissue (Sherman et al., 1971), but rare in plants (Mukherjee and Axt, 1984); detected in trace amounts in the protozoan Tetrahymena vorax (Kersting et al., 2003)

No natural source Occurs in plants such as gymnosperms, a few angiosperms (Dittrich et al., 1971; Dittrich and Kandler, 1972) and seagrass (Drew, 1983), often in methylated form No natural source No natural source

Inositol Phosphates in Soil

OP

PO

PO OP PO OP Fig. 12.4. The structure of muco-inositol hexakisphosphate, which was reported once in plant tissue but never in soil. Phosphate groups are denoted by ‘OP’.

turkeys, dairy and beef cattle) using solution 31P NMR spectroscopy, despite a detection limit of around 1 mg P/kg dry manure (Leytem et al., 2004; Maguire et al., 2004; Turner, 2004). Chemical synthesis is unlikely to be involved in the formation of the phosphorylated stereoisomers in the environment (Cosgrove, 1980). It can occur following extended heating in strong acid or alkali (Cosgrove, 1975) or through simple reactions such as the epimerization of myo-inositol pentakisphosphate to scyllo- and D-chiro-inositol pentakisphosphate by hypobromite oxidation followed by sodium borohydride reduction (Cosgrove, 1972). However, such reactions would clearly not occur in soil. Unphosphorylated inositol stereoisomers, by comparison to the phosphorylated forms, are relatively widespread in nature (Table 12.3). Five are common in plant tissue (myo-, D-chiro-, L-chiro, scyllo- and muco-), often in methylated forms such as pinitol (3-O-methyl D-chiro-inositol) that play a role in stress tolerance (Loewus and Murthy, 2000). A sixth stereoisomer, neo-inositol, occurs in mammalian tissue (Sherman et al., 1971), but has been reported only once in a plant (Mukherjee and Axt, 1984). The remaining three inositols (cis-, epi- and allo-) have no natural source. Only myo-inositol appears to be synthesized directly in plants, with other stereoisomers formed by epimerization reactions (Loewus and Murthy, 2000). Such reactions are similar to the conversion of myo-inositol to scyllo-inositol in bovine brain, which involves the enzymatic epimerization of the C-2 carbon of myo-inositol (Hipps et al., 1973). Synthesis can also occur by cyclization of an appropriate sugar phosphate, as in the formation of neo-inositol 1-phosphate in rat brain via cyclization of mannose 6-phosphate by L-myo-inositol 1-synthase (Sherman et al., 1971).

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Several laboratory studies have detected the formation of inositol phosphates in ‘synthetic’ soils incubated with sugar and nutrients, which suggests a microbial source for the phosphorylated stereoisomers (Table 12.4). Smith and Clark (1951) demonstrated that radiolabelled phosphate added to soil was incorporated into inositol hexakisphosphate after incubation for 30 days with dextrose and ammonium nitrate. Examination of the chromatograms suggests that both myo- and scyllo-inositol hexakisphosphates were present in an approximate 2:1 ratio. This provides unequivocal evidence for the biosynthesis of inositol phosphates by soil organisms, although the specific process leading to incorporation of the labelled phosphate into inositol hexakisphosphates was not resolved (see below). Direct synthesis of inositol phosphates by soil organisms was also demonstrated by incubating ‘synthetic’ soils, or inositol phosphate-free soil, with sugar, inorganic nutrients and a water extract of soil to inoculate the samples with soil microbes (Caldwell and Black, 1958b). Both myo- and scylloinositol hexakisphosphates were subsequently detected, with the scyllo form being more abundant in every sample. Cosgrove (1964) was subsequently able to detect only myo-inositol hexakisphosphate in a similar, albeit shorter, experiment, although it is possible that other phosphorylated stereoisomers were present in undetectable concentrations. Of potential significance is that inositol phosphates were not detected in samples maintained at pH 6, whereas ~3 mg P/kg were detected in two more acidic samples. Important insight into the origin of the phosphorylated stereoisomers was provided by experiments in soil using 14C-labelled compounds (Table 12.4). These demonstrated unequivocally that myo-inositol can be epimerized to D-chiroinositol, and that myo-inositol can be phosphorylated to the hexakisphosphate (L’Annunziata and Gonzalez, 1977; L’Annunziata et al., 1977). The phosphorylation of D-chiro-inositol was not detected, but the study nevertheless provided strong evidence for a microbial source of the phosphorylated stereoisomers in soil. The relative abundance of free inositol isomers in plants raises the possibility that they could be a source of the free isomers in soil. However, the fact that two isomers common in plant tissue (L-chiro- and muco-inositol) do not occur in phosphorylated forms in soil suggests

200

Table 12.4. Studies showing the synthesis of inositol phosphates by soil organisms in laboratory incubations. Reference Smith and Clark (1951)

Soil properties

Nutrients added

Incubation conditions

Inositol phosphates detected

0.5 M NaOH, hypobromite oxidation, ion-exchange chromatography

Radiolabelled myo- and scylloinositol hexakisphosphates (~2:1 ratio) plus some pentakisphosphates

Concentrated HCl, 0.5 M NaOH, ion-exchange chromatography

myo-Inositol hexakisphosphate (0–1.16 mg P/kg) and scyllo inositol hexakisphosphate (0.12–2.14 mg P/kg); scyllo > myo in all samples

1 M NaOH, hypobromite oxidation, ion-exchange chromatography

3.0 mg P/kg myo-inositol hexakisphosphate in the two most acidic samples, none in the pH 6 sample 14 Hot 3 M NaOH, ionC-myo-Inositol exchange chromatography hexakisphosphate, 14C-D-chiroinositol

B.L.Turner

A loam under prairie Dextrose, ammonium Field capacity, room temperature, 30 days (pH 6.7, organic P nitrate and 32P 144 mg P/kg) and a silt loam under forest (pH 5.3, organic P 289 mg P/kg) Caldwell and Various samples of soil Sucrose, inorganic 10 months at 30°C, Black (1958b) C-horizons (calcareous nutrients (also or 3 months at 28°C and acidic), sand and soluble starch in sand–clay mixtures, first experiment) and with pH between soil–water extract 5.1 and 7.3 Cosgrove (1964) Kaolinite and Sucrose, inorganic 5 months, 25°C sand (1:4 ratio) nutrients and soil– maintained at three water extract pH values (4, 5 and 6) 14 L’Annunziata A-horizon of an C-myo -Inositol Field capacity, et al. (1977) uncultivated Andisol 12 days, 36.5°C under forest (pH 5.8, organic matter 11%, clay 18%)

Extraction and analysis

Inositol Phosphates in Soil

that an alternative source is more likely. Of possible significance, therefore, is that the fresh water ciliate Tetrahymena vorax contains the same four inositol stereoisomers in similar proportions to those present in phosphorylated forms in soil (Kersting et al., 2003). Only the free inositols were detected, although a series of higher-order myo-inositol phosphates were identified in Paramecium tetraurelia, another fresh water protozoan (Freund et al., 1992). As protozoa are abundant in soils, they may therefore be an important source of the inositol stereoisomers in free or phosphorylated forms. Despite the evidence for a microbial source of the phosphorylated stereoisomers, no soil organisms have so far been found to contain them. There is no known prokaryotic source of inositol phosphates. Of the eukaryotic microorganisms, Cosgrove (1964) could not detect inositol phosphates in the yeast Saccharomyces carlsbergensis or in 12 fungi isolated from soils rich in phytate. However, S. cerevisiae (Baker’s yeast) contains abundant myo-inositol phosphates, including highly phosphorylated inositol pyrophosphates that appear to phosphorylate proteins (Saiardi et al., 2004). The latter process is selective for eukaryotic proteins, as it was not observed in bacterial extracts.

The Potential Function of Phosphorylated Inositol Stereoisomers in Soil Why do soil organisms synthesize phosphorylated inositol stereoisomers such as scyllo-inositol hexakisphosphate when nature relies almost exclusively on the myo stereoisomer? What are the evolutionary benefits of stereoisomeric inositol phosphates that have favoured their synthesis in soils? And which organisms synthesize them? These are perhaps the most intriguing questions regarding inositol phosphates in soil, but we seem far from answering them. The obvious functional difference between myo-inositol hexakisphosphate and the other phosphorylated stereoisomers that occur in soil is the relative resistance of the latter to enzymatic attack. In particular, scyllo-inositol hexakisphosphate, with no axial groups, seems to be most resistant to phytase activity and is also the most

201

abundant phosphorylated stereoisomer other than myo-inositol found in soil. For example, Greaves et al. (1967) reported that phytase isolated from Aerobacter aerogenes, a Gram-negative facultative anaerobic bacteria, had no activity towards scylloinositol hexakisphosphate. Cosgrove (1970) subsequently found that the ‘SB2’ phytase from a soil Pseudomonas sp. was active towards scyllo-inositol hexakisphosphate, but that the rate of hydrolysis was the slowest of the four isomers tested, being in the order of myo-> neo-> D-chiro-> scyllo-. Other phytases, such as β-propeller phytase and purple acid phytase (see Mullaney and Ullah, Chapter 7, this volume), have not yet been tested against phosphorylated stereoisomers other than myoinositol hexakisphosphate. The resistance of scyllo-inositol hexakisphosphate to hydrolytic attack raises the possibility that it might be synthesized by certain organisms to protect phosphorus from uptake by nearby competing organisms. Such a strategy for conserving phosphorus might be expected to occur in environments where phosphorus is scarce, and could conceivably arise among soil microbes or as part of the complex competition between plants and microbes for soil nutrients (Kaye and Hart, 1997). There is currently no evidence to support this hypothesis, but it warrants investigation. A further possibility is that stereoisomeric inositol phosphates have a non-nutritional function in soils. These highly reactive, but biologically recalcitrant, compounds might, for example, play a role in soil structure (Anderson, 1980) by stabilizing clay–metal–humic complexes in an analogous way to their presence in the core of human RNA-editing enzymes (Macbeth et al., 2005). Alternatively, their capacity to form insoluble precipitates with metals might mean that they are synthesized to ameliorate metal toxicity. For example, the arsenic hyperaccumulating brake fern Pteris vittata secretes inositol hexakisphosphate from its roots (Tu et al., 2004), although it is not known which stereoisomers are involved.

Conclusions and Research Priorities Soils contain large amounts of inositol phosphates, some of which occur nowhere else in nature, yet our understanding of their origin and function in soils is extremely limited. This is unsatisfactory

202

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given the importance of soil phosphorus in the regulation of ecosystems, nourishment of crops and pollution of water bodies. Future research should focus on quantifying the concentrations of the various stereoisomers of inositol hexakisphosphate in soil from a wide range of environments and on investigating their biochemical origin and function. In particular, efforts should be directed towards the following key topics: ●

●

Improvement in analytical procedures for quantification of inositol phosphates in soils. Recent advances in the analysis of soil inositol phosphates by solution 31P NMR spectroscopy mean that the quantification of myo- and scyllo-inositol hexakisphosphate is now straightforward. This should reinvigorate research on these compounds, although continued improvements are of key importance. In particular, the ability to rapidly quantify all four stereoisomeric inositol phosphates that occur in soils is desirable. The use of two-dimensional NMR spectroscopy or mass spectrometry may facilitate this (see Murthy, Chapter 2, and Cooper et al., Chapter 3, this volume). Assessment of the forms and concentrations of inositol phosphates in soil from a wide range of environments. Inositol phosphates are abundant in most mineral soils, but all or none of the soil organic phosphorus can be in this form. More information is therefore required, especially on the phosphorylated stereoisomers, for soil from a variety of environments and ecosystems. Most data are from temperate agroecosystems that are relatively fertile compared to soil under natural vegetation. Little is available for the other major biomes, but this is likely to provide impor-

●

●

tant information on the dynamics and function of the inositol phosphate stereoisomers in soil. Investigation of the dynamics of inositol phosphates in soil. Inositol phosphates are considered to be recalcitrant in soil, yet recent evidence suggests that they are a potentially important source of phosphorus to organisms. In particular, microbes with the capacity to utilize inositol phosphates seem to be widespread in the environment and may be important in making inositol phosphates available to plants. More information is now required on the rates of synthesis and decomposition of inositol phosphates in soil, particularly for the phosphorylated stereoisomers. This will almost certainly involve the use of isotopes (see L’Annunziata, Chapter 4, this volume). Determination of the origins of the phosphorylated inositol stereoisomers in soil and the organisms involved in their synthesis. More than three decades ago, Cosgrove (1972) wrote that ‘[f]inal proof of a microbial origin for the isomers awaits the isolation of soil organisms capable of their biosynthesis’. However, no organism that synthesizes inositol hexakisphosphate stereoisomers other than myo has yet been isolated. The rapid advances being made in DNA-based community analysis should facilitate the investigation of a microbial origin for the stereoisomers, although the possibility remains that other types of organisms, including plants, are also important sources. Confirmation of a biosynthesis pathway for the phosphorylated inositol stereoisomers would be a major advance in our understanding of these enigmatic compounds.

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Magid, J., Tiessen, H. and Condron, L.M. (1996) Dynamics of organic phosphorus in soils under natural and agricultural ecosystems. In: Piccolo, A. (ed.) Humic Substances in Terrestrial Ecosystems. Elsevier Science, Oxford, pp. 429–466. Maguire, R.O., Sims, J.T., Saylor, W.W., Turner, B.L., Angel, R. and Applegate, T. J. (2004) Influence of phytase addition to poultry diets on phosphorus forms and solubility in litters and amended soils. Journal of Environmental Quality 33, 2306–2316. Mahieu, N., Olk, D.C. and Randall, E.W. (2002) Multinuclear magnetic resonance analysis of two humic acid fractions from lowland rice soils. Journal of Environmental Quality 31, 421–430. Martin, J.K. (1970) The measurement of inositol polyphosphates in soil extracts by isotope dilution analysis. New Zealand Journal of Agricultural Research 13, 930–939. Martin, J.K. and Wicken, A.J. (1966) Soil organic phosphorus. IV. Fractionation of organic phosphorus in alkaline soil extracts and the identification of inositol phosphates. New Zealand Journal of Agricultural Research 9, 529–535. Martin, J.-B., Laussmann, T., Bakker-Grunwald, T., Vogel, G. and Klein, G. (2000) neo-Inositol polyphosphates in the amoeba Entamoeba histolytica. The Journal of Biological Chemistry 275, 10134–10140. McGill, W.B. and Cole, C.V. (1981) Comparative aspects of cycling of organic C, N, S and P through organic matter. Geoderma 26, 267–286. McKercher, R.B. and Anderson, G. (1968a) Characterization of inositol penta- and hexaphosphate fractions of a number of Canadian and Scottish soils. Journal of Soil Science 19, 302–310. McKercher, R.B. and Anderson, G. (1968b) Content of inositol penta- and hexaphosphates in some Canadian soils. Journal of Soil Science 19, 47–55. Mehta, N.C., Legg, J.O., Goring, C.A.I. and Black, C.A. (1954) Determination of organic phosphorus in soils. I. Extraction methods. Soil Science Society of America Proceedings 18, 443–449. Morré, D.J., Boss, W.F. and Loweus, F.A. (eds) (1990) Inositol Metabolism in Plants. John Wiley & Sons, New York. Moyer, J.R. and Thomas, R.L. (1970) Organic phosphorus and inositol phosphates in molecular size fractions of a soil organic matter extract. Soil Science Society of America Proceedings 34, 80–83. Mukherjee, R. and Axt, E.M. (1984) Cyclitols from Croton celtidifolius. Phytochemistry 232, 2682–2684. Narasimhan, B., Pliska-Matyshak, G., Kinnard, R., Cartensen, S., Ritter, M.A., von Weymarn, L. and Murthy, P.P.N. (1997) Novel phosphoinositides in barley aleurone cells. Plant Physiology 113, 1385–1393. Omotoso, T.I. and Wild, A. (1970a) Content of inositol phosphates in some English and Nigerian soils. Journal of Soil Science 21, 216–223. Omotoso, T.I. and Wild, A. (1970b) Occurrence of inositol phosphates and other organic phosphate components in an organic complex. Journal of Soil Science 21, 224–232. Posternak, T. (1965) The Cyclitols. Holden Day, San Francisco, California. Raboy, V. (2003) myo-Inositol-1,2,3,4,5,6-hexakisphosphate. Phytochemistry 64, 1033–1043. Richardson, A.E. and Hadobas, P.A. (1997) Soil isolates of Pseudomonas spp. that utilize inositol phosphates. Canadian Journal of Microbiology 43, 509–516. Saiardi, A., Bhandari, R., Resnick, A.C., Snowman, A.M. and Snyder, S.H. (2004) Phosphorylation of proteins by inositol pyrophosphates. Science 306, 2101–2105. Saunders, W.M.H. and Williams, E.G. (1955) Observations on the determination of total organic phosphorus in soils. Journal of Soil Science 6, 254–267. Sherman, W.R., Goodwin, S.L. and Gunnell, K.D. (1971) neo-Inositol in mammalian tissues. Identification, measurement, and enzymic synthesis from mannose 6-phosphate. Biochemistry 10, 3491–3499. Smith, D.H. and Clark, F.E. (1951) Anion-exchange chromatography of inositol phophates from soil. Soil Science 72, 353–360. Stewart, J.W.B. and Tiessen, H. (1987) Dynamics of soil organic phosphorus. Biogeochemistry 4, 41–60. Suzumura, M. and Kamatani, A. (1995) Mineralization of inositol hexaphosphate in aerobic and anaerobic marine-sediments – implications for the phosphorus cycle. Geochimica et Cosmochimica Acta 59, 1021–1026. Thomas, R.L. and Lynch, D.L. (1960) Quantitative fractionation of organic phosphorus compounds in some Alberta soils. Canadian Journal of Soil Science 40, 113–120. Tiessen, H., Stewart, J.W.B. and Moir, J.O. (1983) Changes in organic and inorganic phosphorus composition of two grassland soils and their particle size fractions during 60–90 years of cultivation. Journal of Soil Science 34, 815–823. Tu, S., Ma, L. and Luongon, T. (2004) Root exudates and arsenic accumulation in arsenic hyperaccumulating Pteris vittata and non-hyperaccumulating Nephrolepis exaltata. Plant and Soil 258, 9–19. Turner, B.L. (2004) Optimizing phosphorus characterization in animal manures by solution phosphorus-31 nuclear magnetic resonance spectroscopy. Journal of Environmental Quality 33, 757–766.

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Turner, B.L. (2006) Organic phosphorus in Madagascan rice soils. Geoderma (in press). Turner, B.L. and Newman, S. (2005) Phosphorus cycling in wetlands: the importance of phosphate diesters. Journal of Environmental Quality 34, 1921–1929. Turner, B.L. and Richardson, A.E. (2004) Identification of scyllo-inositol phosphates in soils by solution phosphorus-31 nuclear magnetic resonance spectroscopy. Soil Science Society of America Journal 68, 802–808. Turner, B.L., Papházy, M.J., Haygarth, P.M. and McKelvie, I.D. (2002) Inositol phosphates in the environment. Philosophical Transactions of the Royal Society, London, Series B 357, 449–469. Turner, B.L., Cade-Menun, B.J. and Westermann, D.T. (2003a) Organic phosphorus composition and potential bioavailability in semi-arid arable soils of the western United States. Soil Science Society of America Journal 67, 1168–1179. Turner, B.L., Mahieu, N. and Condron, L.M. (2003b) Quantification of myo-inositol hexakisphosphate in alkaline soil extracts by solution 31P NMR spectroscopy and spectral deconvolution. Soil Science 168, 469–478. Turner, B.L., Cade-Menun, B.J., Condron, L.M. and Newman, S. (2005a) Extraction of soil organic phosphorus. Talanta 66, 294–306. Turner, B.L., Mahieu, N., Condron, L.M. and Chen, C.R. (2005b) Quantification and bioavailability of scylloinositol hexakisphosphate in pasture soils. Soil Biology and Biochemistry 37, 2155–2158. Turner, B.L., Newman, S. and Newman, J. (2006) Organic phosphorus sequestration in subtropical treatment wetlands. Environmental Science and Technology 40, 727–733. Unno, Y., Okubo, K., Wasaki, J., Shinano, T. and Osaki, M. (2005) Plant growth promotion abilities and microscale bacterial dynamics in the rhizosphere of Lupin analyzed by phytate utilization ability. Environmental Microbiology 7, 396–404. Walker, L.R. and del Moral, R. (2003) Primary Succession and Ecosystem Rehabilitation. Cambridge University Press, New York. Williams, C.H. and Anderson, G. (1968) Inositol phosphates in some Australian soils. Australian Journal of Soil Research 6, 121–130. Wrenshall, C.L. and Dyer, W.J. (1941) Organic phosphorus in soils. II. The nature of the organic phosphorus compounds. A. Nucleic acid derivatives. B. Phytin. Soil Science 51, 235–248. Yoshida, R.K. (1940) Studies on organic phosphorus compounds in soil: isolation of inositol. Soil Science 50, 81–89.

13

Abiotic Reactions of Inositol Phosphates in Soil Luisella Celi and Elisabetta Barberis

University of Turin, DIVAPRA Chimica Agraria, via Leonardo da Vinci 44, Grugliasco, 10095 Torino, Italy

Although biotic reactions are the main processes governing the transformation of organic matter in biologically active soils and sediments, abiotic reactions can indirectly affect the fate of organic compounds and control their persistence in the environment. The accumulation of organic phosphorus in soil, often reaching 80% or more of the total phosphorus (Anderson, 1980), is attributed to a series of abiotic processes that hamper the biodegradation of certain compounds (Anderson, 1980; Stewart and Tiessen, 1987; Condron et al., 1990). These processes are related mainly to the high affinity of organic phosphorus for soil mineral colloids, which have a large surface area and a large capacity to retain anions (Tiessen et al., 1983; Guzel and Ibrikci, 1994). Complexation and precipitation with polyvalent cations may also enhance the retention of organic phosphorus in the colloidal phase. This explains why organic phosphorus generally accumulates in the finest soil fractions (Fig. 13.1), with the clay fraction often containing more organic phosphorus than silt (Gburek et al., 2005). Sorption and precipitation with soil cations limit in particular the degradation of phosphate monoesters compared with phosphate diesters and other organic phosphorus compounds. The latter are less strongly stabilized and more likely to be found in soil solution, where they are readily degraded by biological processes. This leads to the accumulation of phosphate monoesters in

soils (Table 13.1), although the distribution of organic phosphorus compounds in the living organisms from where they originate is quite different (Magid et al., 1996). In most soils the inositol phosphates are the most abundant group of phosphate monoesters; they occur in various degrees of phosphorylation (from inositol hexakisphosphate to inositol monophosphate) and up to four stereoisomeric configurations (myo-, scyllo-, neo-, D-chiro-) (Cosgrove, 1980; Harrison, 1987; Turner et al., 2002). The amounts and forms of inositol phosphates in soils are reviewed elsewhere in this volume (see Turner, Chapter 12). This chapter examines the main abiotic processes involved in the stabilization of inositol phosphates in soil. These include adsorption and desorption from soil components, complexation reactions and precipitation with polyvalent cations. The influence of soil solution chemistry on these reactions is also discussed, as well as the effects of inositol phosphates on soil surface properties.

Adsorption of Inositol Phosphates in Soil Phosphate adsorption is one of the most widely studied reactions in soil, whereas adsorption of organic phosphorus compounds has received

©CAB International 2007. Inositol Phosphates: Linking Agriculture and the Environment (eds B.L. Turner, A.E. Richardson and E.J. Mullaney)

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Organic P enrichment ratio

8.0

6.0

4.0

2.0

0.0 0

10

20

30

40

50

Clay (%) Fig. 13.1. Relationship between the organic phosphorus enrichment ratio and the content of clay in different types of soil. (From Gburek et al., 2005.)

much less attention. From a theoretical point of view, the term sorption is used instead of adsorption, because it covers any process that removes reactant from the solution, including adsorption and precipitation (Barrow, 1985, 1993). The extent of adsorption of inositol phosphate is controlled by its concentration in solution, whereas the extent of precipitation is determined by the solubility product of the least soluble inositol phosphate compound. This in turn controls the anion concentration in solution, so the extent of inositol phosphate adsorption is also linked to the nature and concentration of polycations in the system that can form insoluble salts. Moreover, the nature of the inositol phosphates (i.e. the number of phosphate groups and the stereochemical configuration) can further affect both the arrangement of adsorbates on the active surface and the formation of precipitates. Adsorption is also governed by the characteristics of the adsorbate. Soil properties such as pH, mineral composition and texture (Fig. 13.2) can strongly affect the rate and extent of inositol

phosphate adsorption (Anderson et al., 1974; McKercher and Anderson, 1989; Leytem et al., 2002). The greater affinity of some soil components towards these organic compounds is related to their specific surface and porosity, degree of crystallinity and surface charge, and is strongly controlled by the pH of the system.

The role of iron and aluminium oxides In acid soils the sorption of inositol phosphates is reported to be dependent on the contents of amorphous iron and aluminium oxides (Anderson et al., 1974; Harrison, 1987; Pant et al., 1994). Among the different iron oxides, the amorphous types such as ferrihydrite alone, or in association with kaolinite, show a greater capacity to retain myo-inositol hexakisphosphate (Table 13.2) compared to the more crystalline goethite or haematite (Ognalaga et al., 1994; Celi et al., 1999, 2003). The presence of aluminium in the

Table 13.1. Distribution of organic phosphorus fractions (as % of total phosphorus) in soils and growing organisms. (From Magid et al., 1996.)

Nucleic acids Phospholipids Phosphate Monoesters

Escherichia coli

Fungi

Nicotiana

Soils

65 15 20

58 20 22

52 23 25

2 5 50

Abiotic Reactions in Soil

209

2000 myo-IP6 sandy myo-IP6 sandy clay myo-IP6 sandy loam myo-IP6 clay Pi sandy Pi sandy clay Pi sandy loam Pi clay

Sorbed P (µg/g)

1600

1200

800

400

0 0

20

40

60

80

100

Equilibrium P (mg P/m) Fig. 13.2. Isotherms of adsorption of myo-inositol hexakisphosphate (myo-IP6) and inorganic phosphate (Pi) in different types of soil. (Adapted from McKercher and Anderson, 1989; Leytem et al., 2002.)

goethite structure does not affect the extent of adsorption (Cannoni et al., 2004). With the exception of ferrihydrite, the iron oxides show a higher affinity for inositol phosphates than for other organic and inorganic phosphates, as deduced by Langmuir K values (Table 13.2). Adsorption of inositol phosphates occurs through their phosphate groups, which react with iron oxide in the same way as the free phosphate ion via a ligand exchange with the H2O and–OH groups of the surfaces (Parfitt et al., 1976; Goldberg and Sposito, 1985). Evidence of this has been obtained by a combination of quantitative, electrochemical and spectroscopic studies. The different sorption ratios observed between myo-inositol hexakisphosphate and phosphate on the various oxides indicate the involvement of a variable number of phosphate groups in the formation of the bonding and a different arrangement of the molecule depending on the characteristics of the surface (Celi et al., 1999, 2001a, 2003). The number of active sites, the distances between contiguous hydroxyls and the roughness of the mineral surface are the main factors affecting the phosphorus–mineral complex configuration. The phosphate groups that do not react with the surface remain free and make the surface highly negative. The electrical potential of the new surfaces formed by the phos-

phorus–iron oxide complex is related to the extent of adsorption and to the number of phosphate groups that remain free after adsorption (Celi et al., 1999). Fourier-transform infrared spectroscopy shows changes in the P=O and P–O bands of myo-inositol hexakisphosphate after adsorption on iron oxides (Fig. 13.3), due to the formation of Fe–O–P bonds (Celi et al., 1999). The higher electron-donor effect of the O–Fe compared with the –OH group caused a repulsion of electrons to oxygen to form Pδ+–Oδ−, lowering the bond energy of P= Ο and increasing that of P–O (Socrates, 1980). From these results, Ognalaga et al. (1994) and Celi et al. (1999) suggested that adsorption of myoinositol hexakisphosphate on goethite occurred through four of the six phosphate groups, with the remaining two groups being free. On ferrihydrite only two phosphate groups were involved, due probably to the roughness of the oxide preventing the optimal arrangement on the surface (Celi et al., 2003). Two phosphate groups per molecule of inositol phosphate are also bound to the surface of haematite, due in this case to an unfavourable distance between the –OH sites for an optimal arrangement of the molecule on the less reactive surface (L. Celi et al., unpublished data). If iron oxides are associated with other soil components, such as kaolinite, the surface area

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Table 13.2. Langmuir coefficients of adsorption isotherms of myo-inositol hexakisphosphate (myo-IP6) and phosphate on iron and aluminium phyllosilicates, and ferrihydrite–kaolinite systems (Fh–KGa2) at pH 4.5 in 0.01 M KCl. (From Celi et al., 1999, 2003; Cannoni et al., 2004.) Xmax is the maximum amount adsorbed on the minerals while K is the Langmuir constant and indicates the affinity of the adsorbate for the surface. xmax (µmol/m2)a Goethite Ferrihydrite Haematite Bayerite Illite Kaolinite (Fh–KGa2)

myo-IP6 Phosphate myo-IP6 Phosphate myo-IP6 Phosphate myo-IP6 Phosphate myo-IP6 Phosphate myo-IP6 Phosphate myo-IP6 Phosphate

0.64 2.4 2.12 4.57 0.40 0.82 0.40 1.3 0.38 1.0 0.27 0.79 2.24 2.96

(3.8) (12.7) (2.4) (2.4) (2.3) (1.6) (13.4)

K (l/mol)

r 2 (n = 10)

8.0⫻103 5.6⫻103 2.4⫻105 3.2⫻105 6.2⫻104 4.0⫻104 1.3⫻105 7.7⫻104 1.0⫻105 7.5⫻103 1.0⫻105 1.6⫻103 2.2⫻106 9.1⫻104

0.978 0.996 0.994 0.996 0.993 0.994 0.993 0.994 0.993 0.994 0.999 0.923 0.993 0.991

a

The values in parentheses indicate the amount of adsorbed myo-inositol hexakisphosphate expressed as moles of phosphorus.

and porosity are closer to that of the phyllosilicate, while the electrical charge is governed by the iron oxide (Celi et al., 2001b). Adsorption of inositol phosphate is similar to that of ferrihydrite in quantitative terms, despite these different mineralogical characteristics, probably due to a different arrangement of the molecule. On the mixed system inositol hexaphosphate is adsorbed probably through only one of its six phosphate groups (Celi et al., 2003). The affinity of inositol phosphates for oxides seems to be related to the number of phosphate groups involved in the bonding: the higher the number of phosphate groups bound to the surface, the lower the affinity (see Langmuir constants, Table 13.2) as a result of the higher energy necessary to form the multiple bonding and to overcome the conformational hindrance determined by the organic moiety (Celi et al., 1999, 2003). A relatively strong correlation (r2 = 0.877; n = 5) was found between the Langmuir K values for goethite, kaolinite, illite, ferrihydrite and mixed ferrihydrite–kaolinite systems, and the inositol hexakisphosphate/inorganic phosphate maximum molar ratio (Fig. 13.4; Celi et al., 2003). In acidic soils, aluminium oxides can also promote the retention of organic phosphorus (Anderson et al., 1974), although to a lower extent

than iron oxides. The capacity of amorphous aluminium oxides to remove myo-inositol hexakisphosphate from solution (Shang et al., 1990, 1992) was much greater than that of bohemite (Anderson et al., 1974) and bayerite (Cannoni et al., 2004), due to the larger surface area. Anderson and Arlidge (1962) observed a limited capacity of gibbsite to adsorb inositol phosphates. myo-Inositol hexakisphosphate has a higher chemical affinity to the aluminium oxide surface compared to myo-inositol monophosphate, confirming that the interaction is regulated by the functionality of the phosphate groups (Shang et al., 1990, 1992). For instance, three phosphate groups per molecule of myo-inositol hexakisphosphate were involved in bonding to bayerite. This results in a high thermodynamic stability of myo-inositol hexakisphosphate–aluminium oxide complexes and a low activation energy with a higher rate constant. As for inorganic phosphate, adsorption initially proceeds due to the high concentration of adsorbate in the surrounding particle, although studies of this topic are limited. As surface coverage proceeds, the adsorbed myo-inositol hexakisphosphate can impede the approach of other molecules to the surface by imposing negative electrical and steric effects (Shang et al., 1990). As for sorption to iron oxides, desorption

Abiotic Reactions in Soil

211

889 794 Goethite 3119

640

890 794 639 3123

Absorbance

Gt−myo-IP6

1038 629 897 792

3174 1646

Gt−myo-IP6 minus Gt

1078 991 1125 1223

myo -IP6 1642 1385

3372 2927 3397

4000

3000

2000

1500

798 829

1000

498 433

400

Wave number (1/cm) Fig. 13.3. Fourier-transform infrared spectra of goethite (Gt), myo-inositol hexakisphosphate (myo-IP6), the complex formed between myo-inositol hexakisphosphate and goethite (Gt–myo-IP6) and the difference spectrum obtained by subtracting the spectrum of Gt from that of the Gt–myo-IP6 complex (Gt–myo-IP6 minus Gt). (From Celi et al., 1999.)

from aluminium precipitates is limited, and neither plants nor microorganisms are able to release the phosphate groups that remain free after adsorption (Shang et al., 1996), indicating that phosphorus bioavailability and biodegradability are strongly limited by the stability of phosphate surface complexes.

The role of calcium carbonate, clays and organic matter In neutral and basic soils the sorption of inositol phosphates is governed by calcite, clays and organic matter (McKercher and Anderson, 1989). Calcareous soils are those with the ability to immobilize large amounts of phosphate, due to the importance of calcium carbonate on phos-

phate chemistry (Ryan et al., 1984). Calcite can retain myo-inositol hexakisphosphate from solution in amounts that largely exceed the maximum coordination capacity of the surface (Celi et al., 2000). This occurs because, in addition to adsorption on the reactive surface, inositol phosphate can complex the calcium ion in equilibrium with the mineral and form two soluble calcium phytate species, Ca1-phytate and Ca2-phytate, even at a very low concentration of phosphorus, whereas the Ca3–phytate complex precipitates at any pH (Table 13.3; Graf, 1983). Calcium complexation can favour the further dissolution of calcite and hence enhance precipitation of Ca3-phytate. This can lead to the accumulation of organic phosphorus in soil dependent on CaCO3-specific surface area, which is related to the particle size rather than to the total amount of CaCO3 (Holford and Mattingly, 1975; Amer et al., 1985).

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4.0 3 106

KL (I/mol)

3.0 3 106 Fh−KGa2 2.0 3 106

1.0 3 106

Illite Kaolinite Goethite

0.00

0.20

0.40

Ferrihydrite

0.60

0.80

myo -Inositol hexakisphosphate/phosphate ratio Fig. 13.4. Relationship between the Langmuir constants (KL) of goethite, kaolinite, illite, ferrihydrite and ferrihydrite–kaolinite systems (Fh–KGa2) vs. the myo-inositol hexakisphosphate/phosphate molar ratio. (From Celi et al., 2003.)

Clay minerals can also be responsible for the retention of inositol phosphates in neutral or basic soils, although their sorption capacity is less with respect to iron and aluminium oxides. Montmorillonite, and to a much smaller extent bentonite, illite and kaolinite, can adsorb myo-inositol hexakisphosphate (Goring and Bartholomew, 1950; Celi et al., 1999). At pH 4.5, illite adsorbs more than kaolinite (0.38 and 0.27 µmol/m2, respectively), while both phyllosilicates show a higher affinity (>KL) than for phosphate, with a better fitting of the Langmuir model even at high adsorbate concentrations (Celi et al., 1999). This could be due to the fact that the

Table 13.3. Values of the apparent association constants (1/mM) determined at 20°C in 50 mM pH buffer, containing myo-inositol hexakisphosphate at a concentration of 3.04 mM (pH 4.8, 6.0, 7.2) and 0.15 mM (pH 8.4, 9.4, 10.4). (From Graf, 1983.) pH 4.8 6.0 7.2 8.4 9.4 10.4

K1

K2

K3

2.89 12.1 22.7 >1000 >1000 >1000

0.34 2.5 22.7 >1000 >1000 >1000

0.20 0.60 22.7 >1000 >1000 >1000

steric hindrance of myo-inositol hexakisphosphate can prevent the disruption of the two minerals and the formation of aluminium-phosphate salts, in contrast to the phosphate ion, which can displace silicon from the clay structures (Rajan, 1975) and then progressively change the sites available for adsorption, hampering the attainment of true equilibrium. The adsorption mechanism, as hypothesized from the myo-inositol hexakisphosphate/phosphate ratio, suggests that three phosphate groups interact with the surface of both illite and kaolinite, while the other three phosphate groups remain free. However, it is possible that the occupation of adsorption sites is partly hindered by the organic moiety of the organic phosphate. Possibly, if the phyllosilicates are coagulated in an edge-to-face structure, myoinositol hexakisphosphate adsorption is hampered and the number of phosphate groups that remain free is greater than expected. Finally, organic matter in both humic and non-humic forms can participate in the retention of inositol phosphates in soil (Hong and Yamane, 1980, 1981; Borie et al., 1989; Makarov et al., 1997). This can occur through physical or chemical incorporation in the organic matter fraction, direct adsorption on the organic surfaces or indirect adsorption through polyvalent cations that act as bridges to form ternary organic matter–metal– inositol phosphate complexes. Incor-

Abiotic Reactions in Soil

poration of inositol phosphates into the organic matter structure by the formation of covalent bonds has also been hypothesized (Brannon and Sommers, 1985), although this requires further investigation.

Complexation, Precipitation and Mineral Dissolution In addition to adsorption onto solid surfaces, the stabilization of inositol phosphates in soil can be related to their high capacity to complex metal cations (Cosgrove, 1980; Nolan and Duffin, 1987). The order of stability of complexes with myo-inositol hexakisphosphate is copper(II) > zinc > nickel(II) > cobalt(II) > manganese(II) > iron(III) > calcium (Martin and Evans, 1987; Nolan and Duffin, 1987). These complexes can have many stability constants and become soluble at low concentrations or in certain pH ranges (Table 13.3), but as the concentration and pH increase, they become less soluble and can precipitate as insoluble salts (Martin and Evans, 1987). Calcium phytate precipitation as a function of pH and phytate activity occurs even at low pH and can be responsible for a considerable proportion of the loss of inositol phosphates from solution (Celi et al., 2001a). The ability of inositol phosphates to chelate cations can have important consequences for soil processes, because it affects the extent of sorption, changes the speciation of inositol phosphates and the relative composition of organic phosphorus in soil, enhances mineral weathering and may have environmental implications. For instance, interaction of inositol phosphates with iron(III) was reported to transform labile organic phosphorus in manure applied to paddy soils into more resistant forms, due to formation of insoluble iron-phytate (Zhang et al., 1994). The high stability of the cation–inositol phosphate complexes can cause the dissolution of minerals by detaching metals from the surfaces. The formation of insoluble salts can further enhance this process by removing the metal ion from the reaction equilibrium. In fact, the formation of calcium complexes can cause dissolution of calcite (Celi et al., 2000), and the desorption of inositol phosphates from ferrihydrite–kaolinite mixed systems and, to a lesser extent, from ferrihydrite and

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goethite is followed by a consistent release of iron (Celi et al., 2003; Martin et al., 2004). The ability of inositol phosphates to complex metals has received great attention in the medical and biological fields, because the anion can cause metal deficiency, especially in animals with diets rich in seeds (Maga, 1982; Frossard et al., 2000, see Raboy, Chapter 8, this volume). Inositol phosphates have been used in an immobilized form, bound to polyvinylpyridine, for removing heavy metal ions from the solution, thus offering a potential mechanism for decontaminating industrial or mining waste waters (Tsao et al., 1997).

Desorption of Inositol Phosphates from Soil Once sorbed in soil, inositol phosphates are not readily released back to solution. Desorption of myo-inositol hexakisphosphate from iron oxides is a slow reaction that is affected by solution pH (Cabrera et al., 1981; Celi et al., 2003; Martin et al., 2004) and by the degree of phosphorus saturation (Parfitt, 1979; He et al., 1991, 1994; Martin et al., 2002). The configuration of the phosphorus–mineral complex and the formation of the multiple site-bindings play an important role in the strength of the bond and can further reduce the extent of desorption. Thus, no release of inositol phosphate was observed from goethite (Martin et al., 2004), it was negligible from ferrihydrite, whereas it reached 16% of the adsorbed amount at basic pH from ferrihydrite–kaolinite mixed systems (Fig. 13.5; Celi et al., 2003). Desorption of inositol phosphates bound to iron(III) oxides could increase under anaerobic conditions following reduction to iron(II) (Schwertmann, 1991). This may explain the rapid decomposition of inositol phosphates in anaerobic marine sediments (Suzumura and Kamatani, 1995) and the absence of inositol phosphates in wetland soils subjected to anaerobic conditions for most of the year (Turner and Newman, 2005). However, it should be considered that inositol phosphates could re-precipitate with reactive amorphous iron oxyhydroxides, as observed for inorganic phosphate (Sah et al., 1989). Moreover, it was suggested that the reduction of Fe(OOH)-phytate resulted in the formation of iron complexes and then insoluble Fe4phytate (De Groot and Golterman, 1993).

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30 pH 3.5 pH 4.5 pH 7.0 pH 8.5

Desorbed P (%)

25

20

15

10

5

0 myo-IP6

Pi

myo-IP6

Pi

myo-IP6

Pi

Ferrihydrite Fh−KGa2 Goethite Fig. 13.5. Desorption of myo-inositol hexakisphosphate (myo-IP6) and phosphate (Pi) from ferrihydrite, ferrihydrite–kaolinite systems (Fh–KGa2) and goethite, as affected by pH. (Adapted from Celi et al., 2003; Martin et al., 2004.)

Desorption of inositol phosphates from iron oxides is reported to be also affected by the presence of competing ligands such as phosphate, citrate, oxalate and carbonate (Nagarajah et al., 1968; He et al., 1991; Presta et al., 2000; Martin et al., 2004). Other factors, such as the time for desorption, solution/soil ratio and temperature, influence desorption of inorganic phosphate from soils and minerals (Barrow and Shaw, 1975; Barrow, 1983), but have not been systematically investigated for inositol phosphates.

Effects of Solution Characteristics on Abiotic Processes The interaction of myo-inositol hexakisphosphate in soil involves adsorbents with variable charge surfaces and an adsorbate with 12 ionizable –OH groups. The pKa values are reported in Table 13.4. The process is therefore affected by the characteristics of the soil solution, including pH, ionic strength, the nature and concentration of electrolytes and the presence of competing anions.

Solution pH affects inositol phosphate adsorption by influencing both the charge of the reacting surfaces (Barrow et al., 1980; Bolan et al., 1986; Barrow, 1993) and that of the adsorbate, with a change in the relative concentrations of the anionic forms (Fig. 13.6). As the negative charge on both adsorbate and adsorbent tends to increase with pH, there is a reduction in the extent of myo-inositol hexakisphosphate sorption at high pH in soils and on minerals with variable charge surfaces (Anderson and Arlidge, 1962; Anderson et al., 1974; Shang et al., 1992; Celi et al., 2001a). Although HPO42− expresses a greater affinity than H2PO4−, and similarly myoinositol hexakisphosphate with eight charges expresses a greater affinity than the form with six charges, the increasing predominance of these more reactive forms with increasing pH appears insufficient to overcome the repulsive forces raised by the increasing negative charge at the absorbate surface. At low pH the adsorption of myo-inositol hexakisphosphate on goethite is more pronounced than for phosphate (Fig. 13.7; Celi et al., 2001a), probably due to a different arrangement of the molecule on the oxide surface or the formation of

Abiotic Reactions in Soil

215

Table 13.4. Dissociation acid constants (pK) of myo-inositol hexakisphosphate (myo-IP6) (from Costello et al., 1976) and H3PO4 (from Corbridge, 1985). Molecule

pK1

pK 2,3

pK 4–6

pK 7

pK 8

pK 9

pK 10,11

pK 12

myo-IP6a H3PO4

1.1 2.0

1.5 6.8

1.8 12.3

5.7 −

6.9 −

7.6 −

10.0 −

12.0 −

a

myo-Inositol hexakisphosphate = C6H6(H2PO4)6.

insoluble iron-phytate salts if the pH is low enough to cause mineral dissolution (Anderson and Arlidge, 1962; Anderson et al., 1974). With increasing pH the behaviour is opposite, with a more pronounced decrease in adsorption capacity for the organic compound (Anderson et al., 1974; Shang et al., 1992; Celi et al., 2001a). This is attributed to the reduced capacity of myo-inositol hexakisphosphate compared to phosphate to neutralize the hydroxyl ions released from the surface during adsorption (Table 13.4; Shang et al., 1990; Celi et al., 2001a). The decrease in adsorption with increasing pH can facilitate mineralization of inositol phosphates in soil at near-neutral pH (Dalal, 1977), which could account for the greater accumulation of inositol phosphates in acid rather than

alkaline soils (Turner et al., 2002; see Turner, Chapter 12, this volume). This is also related to the optimal conditions for microbial activity, although phytase activity is optimum nearer to pH 5 (Ullah and Gibson, 1987). The sorption of phytase in soils is discussed elsewhere in this volume (see George et al., Chapter 14). In addition to pH, the electrical charge of the adsorbent is affected by the nature and concentration of electrolytes concentrating in the double layer surrounding the charged particles (van Olphen, 1977; Barrow et al., 1980; Bowden et al., 1980; Barrow, 1993). With monovalent cations the adsorption is only slightly affected by the concentration of electrolytes. As this concentration increases, the adsorption of inositol phosphates should decrease at pH values lower than

120

Concentration (%)

100

80

HPO42−

60

H2PO4− IHP8−

40

IHP6− 20

0 2

3

4

5

6

7

8

9

10

pH Fig. 13.6. Aqueous speciation of myo-inositol hexakisphosphate (IHP) and inorganic phosphate at increasing pH. (From Celi et al., 2001a.)

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5.0

Qa (µmol P/m2)

4.0

3.0

2.0

1.0

0.0 2

4

6

8

10

pH Fig. 13.7. Effect of the pH on sorption (Qa) of myo-inositol hexakisphosphate (shaded symbols) and phosphate (open symbols) by goethite in 0.01 M KCl. (From Celi et al., 2001a.)

the point of zero charge of the mineral and increase at pH values higher than the point of zero charge. This would be caused by a reduction of the absolute value of the electric potential at the shear plane, due to compression of the double layer, although no studies have directly addressed this. Surface charge is strongly affected by the presence of polyvalent cations. For instance in the presence of calcium, the surface of goethite remained positive even at high pH (9–10) and in the presence of low concentrations of electrolyte (Celi et al., 1998). The ability of inositol phosphates to form complexes with polyelectrolytes could further favour the interaction with adsorbates by the formation of bridges or salts that precipitate on reacting surfaces. With calcium, the adsorption of myo-inositol hexakisphosphate increases above pH 5, well beyond its capacity to form a monolayer on goethite, due to the simultaneous occurrence of adsorption and precipitation of insoluble calcium-phytates (Celi et al., 2001a). The adsorption of inositol phosphates on soil components is so strong that the competition of other ligands in the bulk solution for the same sites of adsorption is expressed slightly, as observed with phosphate, citrate or carbonate (Anderson et al., 1974; Presta et al., 2000; Martin et al., 2004). Conversely, myo-inositol hexakisphos-

phate can displace phosphate from mineral surfaces, either before or during treatment with the latter, and inhibit phosphate adsorption (Anderson et al., 1974; De Groot and Golterman, 1993; Presta et al., 2000). The release of phosphate and organic matter into solution upon myoinositol hexakisphosphate addition, and the inhibition of their re-sorption, have also been observed in soils (Anderson et al., 1974; De Groot and Golterman, 1993; Leytem et al., 2002) and could accelerate phosphorus transfer to water bodies in runoff (see also Leytem and Maguire, Chapter 10, this volume).

Effects of Inositol Phosphate Sorption on Surface Properties The adsorption of high charge-density anions on colloidal particles creates new surfaces with a different charge and electric potential, thus affecting their dispersion/flocculation behaviour. The adsorption of myo-inositol hexakisphosphate on different iron oxides and phyllosilicates can reverse the initial net positive charge of the surfaces, thus increasing particle–particle repulsive forces and colloidal dispersion (Celi et al., 1999, 2003). This is attributable to the phosphate

Abiotic Reactions in Soil

groups of myo-inositol hexakisphosphate that are not involved in the binding mechanism and that have hydroxyl groups that dissociate at pH > 2 (Table 13.4). The new surface will have a higher charge when fewer phosphate groups are bound; thus, phyllosilicate complexes with myo-inositol hexakisphosphate present a larger charge than iron oxide complexes (Celi et al., 1999). The overall net negative charge of the surface is reached with only low concentrations of inositol phosphate and over a large range of pH, whereas with inorganic phosphate the negative charge is obtained only at pH > 5 and with a high percentage of phosphorus coverage. Monovalent cations in the bulk soil solution affect the changes in surface charge only in terms of absolute values, whereas with polyvalent cations the surface charge remains positive due to the formation of mineral–inositol phosphate–cation complexes that counterbalance the effect of the organic anion (Celi et al., 2001a). The ability of inositol phosphates to detach cations from minerals, as shown for iron released from ferrihydrite (Celi et al., 2003), could have important effects on the weathering of surface minerals, although few studies have been devoted to this topic. Moreover, in contrast to phosphate, the relatively large size of the inositol phosphate molecule should preclude its diffusion into the mineral pores through time, allowing a true equilibrium to be reached more rapidly than with inorganic phosphate.

Summary and Recommendations for Future Research Abiotic reactions are the main processes stabilizing inositol phosphates in soil and limiting their degradation by plants and microorganisms. The affinity of these phosphate monoesters for clays and metal oxide surfaces, their ability to form complexes with polyvalent cations and insoluble salts, and their incorporation in organic structures account for the accumulation of inositol phosphates compared to other organic phosphorus compounds in soil. Recent studies have advanced our understanding of the interaction of inositol phosphates with pure or more complex minerals and organic matter, although some aspects remain unknown. In particular, future

217

studies should address the effects of temperature, solution/soil ratio, concentration of electrolytes and stereoisomeric forms of inositol phosphates other than myo-, on the extent and mechanism of inositol phosphate adsorption. Moreover, the role of organic matter should be expanded, and attention paid to the influence of reaction kinetics on the long-term fate of sorbed inositol phosphates. A more comprehensive investigation of the processes regulating inositol phosphate behaviour under anaerobic conditions is also necessary to understand the potential bioavailability of inositol phosphates under changing redox conditions. The dispersal of colloidal particles following inositol phosphate adsorption on minerals has important environmental implications (see Leytem and Maguire, Chapter 10, this volume). Although adsorption immobilizes a large amount of inositol phosphate in soil, there is a great potential for transfer to water bodies in runoff as particulate phosphorus (Turner, 2005). The dispersion caused by adsorption of inositol phosphates can dramatically affect the transport of colloids in soil and may explain the presence of inositol phosphates in the particulate form found in rivers and lakes (McKelvie et al., 1995; Suzumura and Kamatani, 1995). This can contribute to eutrophication, which is currently a major threat to global water quality (Correll, 1998; Turner et al., 2002; see McKelvie, Chapter 16, this volume). Similarly, the ability of inositol phosphates to detach metals from minerals, together with the potential dispersion of particles upon inositol phosphate adsorption, could enhance mineral weathering and clay or metal translocation through the soil profile, with important effects on pedogenic evolution. In the future, integration of research on abiotic and biotic processes should improve our ability to evaluate the availability of inositol phosphates to plants and their transport to water bodies. Studies should also include other soil organic phosphorus compounds. Integrating this information with the large body of literature devoted to inorganic phosphate will enable a comprehensive understanding of the terrestrial phosphorus cycle, and contribute to the development of land management strategies that combine agronomic productivity with sustainable management of the environment.

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References Amer, F., Mahmoud, A.A. and Sabet, V. (1985) Zeta potential and surface area of calcium carbonate as related to phosphate sorption. Soil Science Society of America Journal 49, 1137–1142. Anderson, G. (1980) Assessing organic phosphorus in soils. In: Khasawneh, F.E., Sample, E.C. and Kamprath, E.J. (eds) The Role of Phosphorus in Agriculture. American Society of Agronomy, Madison, Wisconsin, pp. 411–431. Anderson, G. and Arlidge, E.Z. (1962) The adsorption of inositol phosphates and glycerophosphate by soil clays, clay minerals and hydrated sesquioxides in acid media. Journal of Soil Science 13, 216–224. Anderson, G., Williams, E.G. and Moir, J.O. (1974) A comparison of the sorption of inorganic phosphate and inositol hexaphosphate by six acid soils. Journal of Soil Science 25, 51–62. Barrow, N. J. (1983) A mechanistic model for describing the sorption and desorption of phosphate by soil. Journal of Soil Science 34, 733–750. Barrow, N. J. (1985) Reaction of anions and cations with variable-charge soils. Advances in Agronomy 38, 183–230. Barrow, N. J. (1993) Effects of surface heterogeneity on ion adsorption by metal oxides and by soils. Langmuir 9, 2606–2611. Barrow, N. J. and Shaw, T.C. (1975) The slow reactions between soil and anions. 2. Effect of time and temperature on the decrease in phosphate concentration in the soil solution. Soil Science 119, 167–177. Barrow, N. J., Bowden, J.W., Posner, A.M. and Quirk, J.P. (1980) Describing the effects of electrolyte on adsorption of phosphate by variable charge surface. Australian Journal of Soil Research 18, 395–404. Bolan, N.S., Syers, J.K. and Tillman, R.W. (1986) Ionic strength effects on surface charge and adsorption of phosphate and sulphate by soils. Journal of Soil Science 37, 379–388. Borie, F., Zunino, H. and Martinez, L. (1989) Macromolecular-P associations and inositol phosphates in some Chilean volcanic soils of temperate regions. Communications in Soil Science and Plant Analysis 20, 1881–1894. Bowden, J.W., Posner, A.M. and Quirk, J.P. (1980) Adsorption and charging phenomena in variable charge soils. In: Theng, B.K.G. (ed.) Soils with Variable Charge. New Zealand Soil Science Society, Lower Hutt, New Zealand, pp. 147–166. Brannon, C.A. and Sommers, L.B. (1985) Preparation and characterization of model humic polymers containing organic phosphorus. Soil Biology and Biochemistry 17, 213–219. Cabrera, F., De Arambarri, P., Madrid, L. and Toca, C.G. (1981) Desorption of phosphate from iron oxides in relation to equilibrium pH and porosity. Geoderma 26, 203–216. Cannoni, M., Prati, M., Celi, L., Barberis, E. and Violante, A. (2004) Effetto delle proprietà di superficie degli ossidi sulla ritenzione di fosforo organico e inorganico nel suolo. Proceedings of XXII SICA Congress, Perugia, Italy, 21–24 September 2004, p. 9. Celi, L., Presta, M., Ajmone Marsan, F. and Barberis, E. (1998) Effetto del pH e della natura dell’elettrolita sull’interazione tra goethite ed inositolfosfato. Proceedings of XVI SICA Congress, Ravello (SA), Italy, 30 September–2 October 1998, pp. 89–96. Celi, L., Lamacchia, S., Ajmone-Marsan, F. and Barberis, E. (1999) Interaction of inositol hexaphosphate on clays: adsorption and charging phenomena. Soil Science 164, 574–585. Celi, L., Lamacchia, S. and Barberis, E. (2000) Interaction of inositol phosphate with calcite. Nutrient Cycling in Agroecosystems 57, 271–277. Celi, L., Presta, M., Ajmone-Marsan, F. and Barberis, E. (2001a) Effects of pH and electrolyte on inositol hexaphosphate interaction with goethite. Soil Science Society of America Journal 65, 753–760. Celi, L., Recchi, R., De Luca, G. and Barberis, E. (2001b) Caratterizzazione delle proprietà di superficie di sistemi misti caolinite-ossidi di ferro. Proceedings of XIX SICA Congress, Reggio Calabria, Italy, 25–28 September 2001, pp. 29–35. Celi, L., De Luca, G. and Barberis, E. (2003) Effects of interaction of organic and inorganic P with ferrihydrite and kaolinite–iron oxide systems on iron release. Soil Science 168, 479–488. Condron, L.M., Frossard, E., Tiessen, H., Newman, R.H. and Stewart, J.W.B. (1990) Chemical nature of organic phosphorus in cultivated and uncultivated soils under different environmental conditions. Journal of Soil Science 41, 41–50. Corbridge, D.E.C. (1985) Phosphorus: An Outline of Its Chemistry, Biochemistry and Technology. Elsevier, Amsterdam, The Netherlands. Correll, D.L. (1998) The role of phosphorus in the eutrophication of receiving waters: a review. Journal of Environmental Quality 27, 261–266. Cosgrove, D. J. (1980) Inositol Phosphates: Their Chemistry, Biochemistry and Physiology. Elsevier, Amsterdam, The Netherlands.

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Costello, A. J., Glonek, T. and Myers, T.C. (1976) 31P nuclear magnetic resonance – pH titrations of myo-inositol hexaphosphate. Carbohydrate Research 46, 159–171. Dalal, R.C. (1977) Soil organic phosphorus. Advances in Agronomy 29, 83–117. De Groot, C. J. and Golterman, H.L. (1993) On the presence of organic phosphate in some Camargue sediments: evidence for the importance of phytate. Hydrobiologia 252, 117–126. Frossard, E., Bucher, M., Machler, F., Mozafar, A. and Hurrell, R. (2000) Potential for increasing the content and bioavailability of Fe, Zn and Ca in plants for human nutrition. Journal of the Science and Food Agriculture 80, 861–879. Gburek, W. J., Barberis, E., Haygarth, P.M., Kronvang, B. and Stamm, C. (2005) Phosphorus mobility in the landscape. In: Sims, J.T. and Sharpley, A.N. (eds) Phosphorus: Agriculture and the Environment. American Society of Agronomy, Madison, Wisconsin, pp. 941–979. Goldberg, S. and Sposito, G. (1985) On the mechanism of specific phosphate adsorption by hydroxylated mineral surfaces: a review. Communications in Soil Science and Plant Analysis 16, 801–821. Goring, C.A.I. and Bartholomew, W.V. (1950) Microbial products and soil organic matter. III Adsorption of carbohydrate phosphates by clays. Soil Science Society of America Proceedings 15, 189–194. Graf, E. (1983) Calcium binding to phytic acid. Journal of Agricultural and Food Chemistry 31, 851–855. Guzel, N. and Ibrikci, H. (1994) Distribution and fractionation of soil phosphorus in particle-size separates in soils of western Turkey. Communications in Soil Science and Plant Analysis 25, 2945–2958. Harrison, A.F. (1987) Soil Organic Phosphorus: A Review of World Literature. CAB International, Wallingford, UK. He, Z.L., Yuan, K.N., Zhu, X.Z. and Zhang, Q.Z. (1991) Assessing the fixation and availability of sorbed phosphate in soil using an isotopic exchange method. Journal of Soil Science 42, 661–669. He, Z.L., Yang, X., Yuan, K.N. and Zhu, X.Z. (1994) Desorption and plant-availability of phosphate sorbed by some important minerals. Plant and Soil 162, 89–97. Holford, I.C.R. and Mattingly, G.E.G. (1975) The high- and low-energy phosphate adsorbing surfaces in calcareous soils. Journal of Soil Science 26, 407–417. Hong, J.K. and Yamane, I. (1980) Inositol phosphate and inositol in humic acid and fulvic acid fractions extracted by three methods. Soil Science and Plant Nutrition 26, 491–496. Hong, J.K. and Yamane, I. (1981) Distribution of inositol phosphate in the molecular size fractions of humic and fulvic acid fractions. Soil Science and Plant Nutrition 27, 295–303. Leytem, A.B., Mikkelsen, R.L. and Gilliam, J.W. (2002) Sorption of organic phosphorus compounds in Atlantic coastal plain soils. Soil Science 167, 652–658. Maga, J.A. (1982) Phytate: its chemistry, occurrence, food interaction, nutritional significance, and methods of analysis. Journal of Agricultural and Food Chemistry 30, 1–9. Magid, J., Tiessen, H. and Condron, L.M. (1996) Dynamics of organic phosphorus in soils under natural and agricultural ecosystems. In: Piccolo A. (ed.) Humic Substances in Terrestrial Ecosystems. Elsevier, Amsterdam, The Netherlands, pp. 429–466. Makarov, M.I., Malysheva, T.I., Haumaier, L., Alt, H.G. and Zech, W. (1997) The forms of phosphorus in humic and fulvic acids of a toposequence of alpine soils in the northern Caucasus. Geoderma 80, 61–73. Martin, C. J. and Evans, W. J. (1987) Phytic acid: divalent cation interactions. V. Titrimetric, calorimetric, and binding studies with cobalt (II) and nickel (II) and their comparison with other metal ions. Journal of Inorganic Biochemistry 30, 101–119. Martin, M., Celi, L. and Barberis, E. (2002) The influence of the phosphatic saturation of goethite on phosphorus extractability and availability to plants. Communications in Soil Science and Plant Analysis 33, 143–153. Martin, M., Celi, L. and Barberis, E. (2004) Desorption and plant availability of inositol phosphate adsorbed on goethite. Soil Science 169, 115–124. McKelvie, I.D., Hart, B.T., Cardwell, T.J. and Cattrall, R.W. (1995) Use of immobilized 3-phytase and flow-injection for the determination of phosphorus species in natural waters. Analitica Chimica Acta 316, 277–289. McKercher, R.B. and Anderson, G. (1989) Organic phosphate sorption by neutral and basic soils. Communications in Soil Science and Plant Analysis 20, 723–732. Nagarajah, S., Posner, A.M. and Quirk, J.P. (1968) Desorption of phosphate from kaolinite by citrate and bicarbonate. Soil Science Society of America Proceedings 32, 507–510. Nolan, K.B. and Duffin, P.A. (1987) Effects of phytate on mineral bioavailability. In vitro studies on Mg2+, Ca2+, Fe3+, Cu2+ and Zn2+ (also Cd2+) solubilities in the presence of phytate. Journal of the Science and Food Agriculture 40, 79–85. Ognalaga, M., Frossard, E. and Thomas, F. (1994) Glucose-1-phosphate and myo-inositol hexaphosphate adsorption mechanisms on goethite. Soil Science Society of America Journal 58, 332–337. Pant, H.K., Edwards, A.C. and Vaughan, D. (1994) Extraction, molecular fractionation and enzyme degradation of organically associated phosphorus in soil solutions. Biology and Fertility of Soils 17, 196–200.

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Parfitt, R.L. (1979) The availability of P from phosphate–goethite bridging complexes. Desorption and uptake by ryegrass. Plant and Soil 53, 55–65. Parfitt, R.L., Russell, J.D. and Farmer, V.C. (1976) Confirmation of the surface structure of goethite and phosphated goethite. Journal of Chemical Society and Faraday Transaction I 72, 1082–1087. Presta, M., Celi, L. and Barberis, E. (2000) Competizione tra fosfato e inositol fosfato su goethite. Proceedings of XVIII Congress SICA., Catania, Italy, 22–24 September 2000, pp. 100–108. Rajan, S.S.S. (1975) Phosphate adsorption and the displacement of structural silicon in an allophane clay. Journal of Soil Science 26, 250–256. Ryan, J., Curtin, D. and Cheema, M.A. (1984) Significance of iron oxides and calcium carbonate particle size in phosphate sorption by calcareous soils. Soil Science Society of America Journal 48, 74–76. Sah, R.N., Mikkelsen, D.S. and Hafez, A.A. (1989) Phosphorus behavior in flooded–drained soils. II. Iron transformation and phosphorus sorption. Soil Science Society of America Journal 53, 1723–1729. Schwertmann, U. (1991) Solubility and dissolution of iron oxides. Plant and Soil 130, 1–25. Shang, C., Huang, P.M. and Stewart, J.W.B (1990) Kinetics of adsorption of organic and inorganic phosphates by short-range ordered precipitate of aluminium. Canadian Journal of Soil Science 70, 461–470. Shang, C., Stewart, J.W.B. and Huang, P.M. (1992) pH effects on kinetics of adsorption of organic and inorganic phosphates by short-range ordered aluminum and iron precipitates. Geoderma 53, 1–14. Shang, C., Caldwell, D.E., Stewart, J.W.B., Tiessen, H. and Huang, P.M. (1996) Bioavailability of organic and inorganic phosphates adsorbed on short-range ordered aluminium precipitate. Microbial Ecology 31, 29–39. Socrates, G. (1980) Infrared Characteristic Group Frequencies. John Wiley & Sons Chichester, UK. Stewart, J.W.B. and Tiessen, T. (1987) Dynamics of soil organic phosphorus. Biogeochemistry 4, 41–60. Suzumura, M. and Kamatani, A. (1995) Origin and distribution of inositol hexaphosphate in estuarine and coastal sediments. Limnology and Oceanography 40, 1254–1261. Tiessen, H., Stewart, J.W.B. and Moir, J.O. (1983) Changes in organic and inorganic phosphorus composition of two grassland soils and their particle size fractions during 60–90 years of cultivation. Journal of Soil Science 34, 815–823. Tsao, G.T., Zheng, Y.Z., Lu, J. and Gong, C.S. (1997) Adsorption of heavy metal ions by immobilized phytic acid. Applied Biochemistry and Biotechnology 63–65, 731–741. Turner, B.L. (2005) Organic phosphorus transfer from terrestrial to aquatic environments. In: Turner, B.L., Frossard, E. and Baldwin, D.S. (eds) Organic Phosphorus in the Environment. CAB International, Wallingford, UK, pp. 269–294. Turner, B.L. and Newman, S. (2005) Phosphorus cycling in wetland soils: the importance of phosphate diesters. Journal of Environmental Quality 34, 1921–1929. Turner, B.L., Papházy, M.G., Haygarth, P.M. and McKelvie, I.D. (2002) Inositol phosphates in the environment. Philosophical Transactions of the Royal Society, London, Series B 357, 449–469. Ullah, A.H. J. and Gibson, D.M. (1987). Extracellular phytase from Aspergillus ficuum NRRL 3135: purification and characterization. Preparative Biochemistry 17, 63–91. van Olphen, H. (1977) An Introduction to Clay Colloid Chemistry. Wiley-Interscience, London. Zhang, Y.S., Werner, W., Scherer, H.W. and Sun, X. (1994) Effect of organic manure on organic phosphorus fractions in two paddy soils. Biology and Fertilizer Soils 17, 64–68.

14

Interactions Between Phytases and Soil Constituents: Implications for the Hydrolysis of Inositol Phosphates

Timothy S. George1, Hervé Quiquampoix2, Richard J. Simpson3 and Alan E. Richardson3 1

Scottish Crops Research Institute, Invergowrie, Dundee DD2 5DA, UK; 2Unité de Science du Sol, INRA-ENSAM, 2 Place Pierre Viala, 34060 Montpellier Cedex 1, France; 3CSIRO Plant Industry, PO Box 1600, Canberra, ACT 2601, Australia

A large proportion (up to 80%) of soil phosphorus occurs in organic forms (Harrison, 1987), of which derivatives of inositol phosphates constitute a considerable fraction (Anderson, 1980; see Turner, Chapter 12, this volume). The bioavailability of inositol phosphates depends on their mineralization by phytases (myo-inositol-hexakisphosphate phosphatases), which come in several classes (EC 3.1.3.8, EC 3.1.3.26 and EC 3.1.3.72), that initially cleave phosphate at different positions on the myo-inositol ring (see Mullaney and Ullah, Chapter 7, this volume). Phytases were first recognized almost a century ago (Suzuki et al., 1907; Dox and Golden, 1911) and have many biological sources, including plants, animals and a wide range of microorganisms (Table 14.1). They are particularly important in the soil environment, having been identified in plant roots, fungi, yeasts and bacteria (Irving, 1980). Phytases do not constitute a major component of plant root exudates and appear to be absent from monogastric animal digestive systems (Hayes et al., 1999; Brinch-Pedersen et al., 2002). Therefore, research into the role of phytases in biological phosphorus cycling in both natural and agricultural systems has focused primarily on phytases produced by microorganisms. Both soil bacteria and fungi produce extracellular phytases, which

give plants a nutritional benefit when present in the rhizosphere (Findenegg and Nelemans, 1993; Richardson et al., 2001b; Idriss et al., 2002). Microbial phytases have also been specifically engineered for supplementation of monogastric animal feeds (Lehmann et al., 2000) and have the potential to enter soil through animal excreta. In recent years, transgenic plants that express microbial phytase genes have been produced for use in animal diets (Pen et al., 1993) and have been evaluated for their capacity to improve plant nutrition (Richardson et al., 2001a; see Richardson et al., Chapter 15, this volume). Transgenic animals with enhanced phytase activity in saliva have also been developed (Golovan et al., 2001). Of particular significance to the mineralization of inositol phosphates in the soil environment is the expression of fungal (Richardson et al., 2001a; Zimmermann et al., 2003) and bacterial (Lung et al., 2005) phytase genes in plants. These plants show improved phosphorus nutrition when grown under controlled conditions, but this is compromised when plants are grown in the more complex soil environment (George et al., 2004, 2005a,c). Whilst mobility of both substrate (inositol phosphates) and product (phosphate) of the phytase reaction are likely to be major limitations to the efficacy of phytases in the soil environment

©CAB International 2007. Inositol Phosphates: Linking Agriculture and the Environment (eds B.L. Turner, A.E. Richardson and E.J. Mullaney)

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Table 14.1. Biochemical properties of a selection of phytase enzymes from a range of biological sources. (Adapted from Vats and Banerjee, 2004.) Molecular weight (kDa)

Isoelectric point (pl)

pH optimum

Bacillus sp. DS 11 Bacillus subtilis B. licheniformis

44 36–38 44, 47

5.3 6.3 5.0, 5.1

7.0 6.0–6.5 4.5–6.0

70 60 55–65

Escherichia coli

42

6.3–6.5

4.5

60

Kim et al. (1998a,b) Kim et al. (1999) Kerovuo et al. (1998, 2000) Greiner et al. (1993)

120

–

2.0–2.5

55–60

Han et al. (1999)

Aspergillus niger (phyA) A. niger (phyB)

85

4.5

2.5, 5.0

58

Ullah and Gibson (1987)

68

4.0

2.5

63

A. oryzae A. fumigatus Peniophora lycii (phyA)

120 91 72

4.2 7.3 3.6

5.5 4.0–4.5

50 55 50–55

Erlich et al. (1993); Ullah and Cummins (1987) Shimuzu (1993) Wyss et al. (1999b) Lassen et al. (2001)

60

5.5

4.5–4.8

55

Gibson and Ullah (1988)

Source of phytase

Temperature optimum (°C)

Reference

Bacteria

Yeast Saccharomyces cerevisiae Fungi

Plants Glycine max

(see Celi and Barberis, Chapter 13, this volume), edaphic factors directly affecting the stability and catalytic efficiency of phytases are also important (George et al., 2005b, 2006). In this chapter we review the interactions of phytases with soil constituents and consider the implications of this for the hydrolysis of inositol phosphates in soil. Although many edaphic factors alter the synthesis and secretion of phytases through direct impacts on biological systems, this review focuses on impacts of the soil environment on the stability and catalytic efficiency of discrete phytase proteins following their release from the cytoplasm.

Factors Affecting Phytase Activity in Soil Most of the phytases produced by soil organisms are thought to be released as extracellular enzymes by active exudation (Tarafdar et al., 2002) and thus have only a short contact time with the cytoplasmic environment. However, some of the

activity found in soil will presumably be passively released following cell lysis and thus be adapted specifically to the intracellular environment. Once in soil, phytases must withstand many factors in order to remain functional (Fig. 14.1). These include (Goldstein, 1976; Gianfreda and Bollag, 1996; Nannipieri et al., 1996; Nannipieri and Gianfreda, 1998): 1. deactivation and inhibition by adsorption and immobilization on soil solid particles; 2. proteolytic and microbial mediated degradation; 3. inhibition by interaction with metal ions, anions and metabolites; and 4. denaturation by soil environmental factors (temperature, pH, water content, light). The impact of these factors on phytases found in the soil environment is potentially large (Fig. 14.1), leaving little phytase activity for longer-term hydrolysis of inositol phosphates. Few studies have explicitly measured phytase activity in soil, but ‘baseline’ activities, if detectable, are in the range of 10–300 pKat/g soil ( Jackman and Black, 1952;

Sources of phytase input to soil

Interactions Between Phytase and Soil Constituents

223

Endogenous

Plant Proteolysis

Bacteria Fungus Yeast Manures and diet supplement

Soil Plant residues solution

Microbial degradation Adsorption Interaction with ions Soil environment − pH − temperature − water

Inositol phosphates

Soil phytase Soil solid

Microbial

− light

Manures Loss of Phytase activity with time

Fig. 14.1. Sources and fate of phytases in the soil environment. The schematic demonstrates that the biological origin of phytases in soil is varied and that phytase activity may be lost through a wide range of competing biological, chemical and physical processes before its interaction with inositol phosphates, either within the soil solution or at the boundary with the solid phase.

T.S. George et al., 2002 unpublished data), with the maximum represented by a mor humus layer from a spruce woodland (Svenson, 1986). In comparison with the activity of enzymes released to the soil through various biological processes ( Jackman and Black, 1952), these activities are small. For example, samples taken from the rhizosphere of transgenic plants, which exude large amounts of Aspergillus niger phytase, had activities against inositol phosphate representing less than 1% of that known to be exuded to the soil by the plants (George et al., 2004, 2005b). Moreover, baseline phytase activity in soil appears insignificant when compared with total phosphomonoesterase activities, which are 1–2 orders of magnitude greater, but is similar to those of soil phosphodiesterases (Eivazi and Tabatabai, 1977). In spite of this, tolerance of phosphatases (including phytases) to the extracellular environment will vary depending on biochemical properties of the enzyme (Table 14.1), which will have an important influence on the baseline activity in soil and thus to the biological cycling of inositol phosphates.

Interaction of phytases with the soil solid phase Proteins have an affinity for the interface between the aqueous and solid phase of soil, so

adsorption of enzymes is common (Norde and Lyklema, 1991). In some cases this can inhibit enzyme activity irreversibly (Quiquampoix, 1987a, 2000; Quiquampoix and Mousain, 2005). Adsorption of phytases may reduce the affinity for substrates and thus reduce the effective activity. However, immobilization protects phytases from degradation (Naidja et al., 2000) and may be responsible for their long-term persistence in soil (Nannipieri et al., 1996). Processes of immobilization and adsorption Phytase from A. niger was rapidly sorbed when added to a range of soils with varying adsorption capacity (George et al., 2005b). However, most of the activity was immediately recovered on the soil solid phase. This indicates that the initial fate of this particular phytase upon introduction to soil was immobilization by adsorption to soil solid constituents. This may involve binding to solid supports by covalent bonds, cooperative adsorptive interactions, and entrapment and encapsulation in stable aggregates (Gianfreda and Bollag, 1996). Electrostatic and van der Waals forces, as well as hydrophobic interactions, have been suggested for adsorption to clays (Quiquampoix, 2000; Quiquampoix et al., 2002), and intercalation of proteins in layered clays may also occur. In contrast, ion exchange, entrapment in organic networks and covalent bonds may account

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for the stable association between phytases and humic materials (Perez-Mateos et al., 1991; Gianfreda and Bollag, 1996). In addition to charged amino-acid residues, protein groups that are potentially free to react with soil constituents include carboxyl, phenol, thiol, aliphatic hydroxyl and amide groups (Brown and Hasselberger, 1971). The heterogeneous nature of proteins also means they contain both polar and non-polar domains that can interact with hydrophylic and hydrophobic surfaces in the soil, respectively (Laidler and Sundaram, 1971). Soils are also extremely heterogeneous. They contain a mixture of organic materials of various levels of decomposition, clay and metal (hydr)oxide minerals, and combinations of these materials, which produce complexes with a wide range of adsorption properties. Furthermore, enzymes can be adsorbed to live biological material such as cell walls and mucigel (Ciurli et al., 1996; Marzadori et al., 1998). Extracellular enzymes in soil are primarily associated with high-molecular weight organic matter and claysized particles, including specific clay minerals and humified organic matter (Kanazawa and Filip, 1986; Perez-Mateos and Rad, 1989; Rojo et al., 1990; Marx et al., 2005). However, detailed cytochemical and microscopy techniques have been unable to verify the presence of discrete enzyme–clay complexes (reviewed by Ladd et al., 1996). While coarse organic matter and fine clay fractions are likely to be the main sites for adsorption and immobilization of phytases, they are also the primary habitats for microorganisms, the major source of phytases in soil (Rojo et al., 1990; Marx et al., 2005). Despite the heterogeneity of soil, predictable patterns of biological and biochemical activity are evident. For example, biological activity and its products decline with depth, which has implications for the distribution of phytase activity in terms of its production and the availability of organic sites for immobilization (Nannipieri and Gianfreda, 1998). Moreover, the major zone of concentration of phytases in soil is likely to be the rhizosphere, where plant and soil are in contact and microorganisms are abundant (Tinker and Nye, 2000). As such, the rhizosphere will likely be the initial external environment experienced by many extracellular phytases. Clay minerals will provide a major proportion of the surface area for adsorption of phytases

in many soils. The adsorption capacity of the clay fraction depends on the type of clay present (e.g. layered clays vs. non-layered clays, permanent charge vs. variable charge minerals) with major differences being exemplified by comparisons between montmorillonite and kaolinite, the former being the stronger adsorbate. While much of the theoretical research on adsorption of enzymes by clays has involved pure forms of montmorillonite (Quiquampoix and Mousain, 2005), such model systems may be poorly representative of the soil environment. ‘Perfect clays’ rarely exist in soil, where magnesium, aluminium and silicon in the mineral lattice are commonly substituted for by impurities, causing changes in the electrical charge of the clay (Rowell, 1994). Dispersion of clays by large concentrations of certain cations such as sodium and potassium can also modify sorption surfaces and affect enzyme adsorption (Violante and Gianfreda, 2000). In addition, clays are often coated with organic material or metal (hydr)oxides, which alters their adsorption capacity. For example, enzymes are less adsorbed on aluminium-coated montmorillonite than on pure clay (Violante and Gianfreda, 2000). Even the presence of phytases (and other proteins) may alter the adsorption environment presented by the clay, as rapid unfolding of the enzyme at the clay surface may produce a denatured protein monolayer covering the clay particle, acting as a more benign adsorption environment for subsequent enzymes (Brown and Hasselberger, 1971). In addition to interaction with mineral materials, phytases also form complexes with humic substances and their constituents, including phenols and quinones (Ladd and Butler, 1975; Wetzel, 1993; Gianfreda and Bollag, 1996). Reactions can be reversible (e.g. hydrogen bonding between phenolic groups and oxygen in the peptide bond) or irreversible (e.g. covalent bonding between terminal amino and sulphhydryl groups of the enzyme) (Ladd and Butler, 1975). Further interaction with other soil components may also occur; for example, flocculation of organic–protein complexes was enhanced by the presence of polyvalent cations and clay minerals (Rao and Gianfreda, 2000; Violante and Gianfreda, 2000). Phytases added to soil collected from the rhizosphere of plants were less rapidly adsorbed to the solid phase than when added to soil that had not been affected by the presence of a

Interactions Between Phytase and Soil Constituents

growing plant root (George et al., 2005b). In a soil with relatively little sorption capacity, phytases from A. niger remained active in the solution phase to a greater extent in rhizosphere soil than in bulk soil. In contrast, in soil with greater clay and organic matter content and larger cation–anion exchange capacity, a smaller amount of phytases remained in solution under rhizosphere conditions. Such differences are likely to be associated with differences in the chemistry and biochemistry of rhizosphere and bulk soil, such as pH and the presence of proteins and organic anions. The presence of organic anions, such as citrate and malate, has been shown to increase the amount of acid phosphatase retained in soil solution (Huang et al., 2002, 2003) due to ligand exchange reactions of these anions in the presence of metal (hydr)oxide surfaces (Violante and Gianfreda, 2000). Given the heterogeneous charge characteristics of proteins (including phytases), competitive exchange reactions will involve anions as well, while other hetero-valent proteins may be of similar importance. The significance of electrostatic interactions between phytases and soil constituents will also depend on the ionic strength of the soil solution, as electrostatic forces diminish with increasing salt concentration (Goldstein, 1976). Differences in ionic strength may be especially important in determining retention of phytases where soil moisture fluctuates and salinity and sodicity are apparent. Soil pH exerts a strong control on phytase adsorption. Adsorption of an A. niger phytase to a range of soils was shown to be complete at pH 4.5, close to the isoelectric point of this enzyme (pH 4.8–5.2; Wyss et al., 1999a). In contrast, the proportion of phytase activity recovered in soil solution was greater as pH was increased (Fig. 14.2). A number of enzymes have maximum adsorption on a range of different surfaces at their isoelectric point (Kondo et al., 1993; Quiquampoix et al., 1993, 2002; Violante et al., 1995; Huang et al., 1999). Proteins at their isoelectric point have no net electric charge, and therefore only weak interactions such as van der Waals or hydrogen bonding are invoked for such adsorption (Violante and Gianfreda, 2000). Nevertheless, entropic factors, such as hydrophobic interactions or modification of protein conformation towards a less-ordered secondary structure, can result in stronger interactions with

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soil constituents, even at a protein’s isoelectric point. The general partitioning of phytase activity to the solution phase with increased pH is attributable to greater electrostatic repulsion above the isoelectric point of the protein due to enthalpic forces (Kondo and Higashitani, 1992; Kondo et al., 1993; Quiquampoix et al., 1993). To investigate further the importance of the isoelectric point for adsorption of phytases, the adsorption of Peniophora lycii phytase (isoelectric point ≈ 3.6) and that produced by A. niger (isoelectric point ≈ 4.8) were compared following addition to a range of soils (George et al., 2006). It was shown that P. lycii phytase remained active in solution at a soil pH of ~5.5, whereas A. niger phytase was rapidly adsorbed to soil solid phase (Fig. 14.3). This is potentially important as the known range of isoelectric points (pH 3.6–7.3) for phytases is large (Table 14.1), suggesting that different enzymes will be more or less adsorbed over a wide range of soil pH. Biochemical differences in proteins have also been invoked in studies of the adsorption of phosphatases from ectomycorrhizal fungi, with interspecific and intraspecific differences in adsorption properties being observed for different enzymes (Quiquampoix and Mousain, 2005). Phosphatases from Pisolithus tinctorius showed no adsorption on montmorillonite between pH 2 and 8, whereas phosphatases from other species (Cenococcum geophilum, Hebeloma cylindrosporum) showed an increasing adsorption from pH 6 to 4, followed by complete inhibition of the catalytic activity of the adsorbed fraction. Furthermore, adsorption of phosphatase from Suillus bellini varied with pH, whereas that from S. mediterraneensis was completely adsorbed across a range of pH, although catalytic activity of both enzymes was maintained regardless of their adsorption (Quiquampoix and Mousain, 2005). It is therefore apparent that not only does specific variability in the adsorption of distinct phytases occur, but differences in inhibition by adsorption are also probable. Differences in biochemical characteristics of phytases will affect their mobility and are therefore potentially important for the interaction of phytases with inositol phosphates (Wyss et al., 1999a; Vats and Banerjee, 2004; Quiquampoix and Mousain, 2005; George et al., 2006).

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(i) 100

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8 days Fig. 14.2. The effect of pH on the adsorption of Aspergillus niger phytase in soil. Phytase activity, measured as a proportion (%) of the amount initially added to soil, is shown (i) in the solution phase and (ii) on the solid phase of two soil types: (a) a Spodosol and (b) an Alfisol. Phytase activity against myoinositol hexakisphosphate was measured at three time points (1 h, 48 h and 8 days) after addition of phytase to soil suspensions buffered at a range of pH 4.5–8.5. Data show the mean of three replicates with bars representing two standard errors. Differences between enzyme activity measured at different pH and at different times (within each soil type) for both solution and solid phase were established using ANOVA. Least significant difference (P < 0.05) is presented as a bar for each soil type and phase within soil. (From George et al., 2005b.)

Inhibition and protection of phytases by immobilization Protection of enzymes has been demonstrated upon adsorption to a number of soil surfaces including clay, metal (hydr) oxides, organic material and mucigel produced by plants and microorganisms (Rao et al., 1994; Ciurli et al., 1996; Naidja et al., 2000). Immobilized enzymes usually show increased resistance to temperature, protease activity and microbial degradation (Estermann et al., 1959; Makboul and Ottow, 1979; Sarkar et al., 1989; Kandeler, 1990; Perez-Mateos et al.,

1991; Nannipieri, et al., 1996; Naidja et al., 2000; Violante and Gianfreda, 2000) and improved resistance to freezing and thawing, wetting and drying, changes in pH and presence of heavy metals (Gianfreda and Bollag, 1996). Once adsorbed, A. niger phytases added to a range of soils showed greater stability than enzymes not in the presence of soil (George et al., 2005b, 2006). Moreover, the activity of endogenous soil phosphatases appears to be more stable than newly added or recently immobilized enzymes, presumably due to persistence of

Interactions Between Phytase and Soil Constituents

(a) 12

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10 8 6 4 2 0 0

Peniophora lycii; pH 5.5 Peniophora lycii; pH 7.5 Aspergillus niger; pH 5.5 Aspergillus niger; pH 7.5

Fig. 14.3. Adsorption of phytase from Aspergillus niger and Peniophora lycii as a function of differing isoelectric points (pI). Activity (nKat/g soil) against myo-inositol hexakisphosphate of the two fungal phytases with different pI (4.8 and 3.6, respectively) was measured in the solution phase of two soil types: (a) a Spodosol and (b) an Alfisol. Phytase activity was measured at time points between 1 and 24 h after the addition of phytases to soil suspensions buffered at pH 5.5 and 7.5. Data show the mean of three replicates with bars representing two standard errors. (From George et al., 2006.)

stabilized forms of the enzymes over time (PerezMateos et al., 1991). The resilience of immobilized phytases may be due to a number of factors including: (i) concurrent adsorption of proteases and inhibitory substances (metal ions, chelators, etc.) (Ciurli et al., 1996; Demanèche et al., 2001; Casucci et al., 2003; Renella et al., 2003); (ii) steric hindrance against relatively large proteases and microorganisms for enzymes embedded in organic matrices (Wetzel, 1993); and (iii) confor-

227

mational changes that increase the stability of the protein structure, preventing autolysis and increasing the energy required for denaturation (Quiquampoix and Mousain, 2005). Despite the possibility of protection of a proportion of phytase activity by adsorption, much activity may also be inhibited by interaction with soil solid surfaces (George et al., 2005b). Generally, adsorption in mixed environments such as soil is less inhibitory to enzyme activity than adsorption to pure clays or some organic materials. For example, several studies demonstrated less inhibition of phosphatase activity when adsorbed to montmorillonite coated with aluminium hydroxides than on clean clay surfaces (Rao et al., 1994; Geiger et al. 1998a; Bayan and Eivazi, 1999; Huang et al., 1999). Reduced inhibition has similarly been observed for aluminium hydroxide–tannic acid complexes compared with tannic acid alone (Gianfreda et al., 1993). Soil phosphatase activity is commonly correlated with organic matter content, whereas negative relationships with clay content are often observed (Harrison, 1983; Feller et al., 1994). Faster inhibition of phytase and other phosphatases following adsorption has similarly been shown to occur in soil with increasing dominance of clay compared to organic matter (Sarkar et al., 1989; George et al., 2005b). The temporal pattern of phytase degradation following adsorption to soil solid constituents (George et al., 2005b) was similar to those observed for acid phosphatase adsorbed to various clay–sesquioxide and organic surfaces (Rao et al., 2000) and other enzymes on mixed clay–sesquioxide–organic surfaces (Nannipieri et al., 1996; Naidja et al., 2000). Interestingly, the temporal unfolding of model proteins adsorbed on montmorillonite (Quiquampoix et al., 2002) also follows a similar pattern, suggesting a potential mechanism for phytase deactivation after adsorption to soil (Leprince and Quiquampoix, 1996). This is corroborated by the fact that loss of A. niger phytase activity on the soil solid phase is irreversible (George et al., 2005b). The rate of irreversible inhibition after adsorption is also pH-dependent (Fig. 14.1) and a function of electrostatic forces. At pH below the isoelectric point of the phytase, increased conformational change and denaturation following adsorption would be expected (Leprince and Quiquampoix, 1996).

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The resilience of phytase in the soil environment also depends on the type of clay mineral present; for example, kaolinite is less inhibitory to adsorbed enzymes than clay with interlaminar spaces, such as montmorillonite (Nannipieri et al., 1996). Moreover, the capacity of pure montmorillonite for adsorption and inhibition of phosphatase is greater than intercalated equivalents, suggesting that phosphatases are absorbed and strongly inhibited in the interlayer region of the pure clay (Kelleher et al., 2004). The potential importance of organic matter in protecting phytase activity is implied by the fact that remnant phosphatase activities in soil are associated almost exclusively with organic, rather than clay, surfaces (Ladd et al., 1993). The protective effect of organic material has been attributed to it being structurally diverse, with macropores where enzymes can lodge and be sterically protected from microorgansims and proteolytic enzymes, while still permitting access by relatively low-molecular weight substrates (Estermann et al., 1959; Burns et al., 1972; Naidja et al., 2000). In fact, the larger the humic–enzyme complex is, the greater is the apparent resistance of the immobilized enzyme to degradative factors (Nannipieri et al., 1988). The organic portion of clay–humus complexes is considered to be important in protecting enzymes from the inhibitory effects of clay, by acting as a barrier between clay and enzyme (Quiquampoix, 1987b; Quiquampoix et al., 1995, 2002). However, some organic compounds can also be inhibitory to phosphatase activity. For example, activity is reduced when phosphatase is complexed with tannic acid (Rao and Gianfreda, 2000) and phenolics (Wetzel, 1993), with inhibition being caused by blocking of the enzyme active site (Ladd and Butler, 1975). Paradoxically, phosphatase–tannic acid complexes formed in the presence of both iron-oxides and montmorillonite retain greater activity than in the presence of the tannic acid or minerals alone (Rao and Gianfreda, 2000). Of particular interest is that the resilience of phytase towards the adsorption environment appears to be determined by whether the enzymes usually have extracellular or intracellular function. For example, intracellular phytases from wheat and S. collinitus were completely adsorbed on clay particles across a range of pH, and showed complete inhibition of catalytic activ-

ity (Matumoto-Pintro and Quiquampoix, 1997). In comparison, two extracellular phytases from A. niger and H. cylindrosporum retained significant catalytic activity in the presence of clay, irrespective of their degree of adsorption (MatumotoPintro and Quiquampoix, 1997). Similarly, extracellular phosphatase from maize roots was less inhibited in soil than that from wheat germ (Dick et al., 1983). Such retention of activity by extracellular enzymes upon adsorption has been suggested to be an evolutionary consequence of an organism’s requirement for a functional extracellular enzyme (Quiquampoix, 2000). Indeed, it has been suggested that adsorbed–uninhibited enzymes may act as an indicator of transient concentrations of biochemically available substrates to microorganisms in the surrounding soil niche (Burns, 1982; Allison and Vitousek, 2005). Effects of phytase immobilization on reaction kinetics The heterogeneity of the soil environment means that most extracellular phytase will not catalyse reactions as efficiently as in homogeneous in vitro systems. The kinetics of immobilized phytase in soil are likely to be different to that of free phytase in soil solution (Nannipieri and Gianfreda, 1998). Studies with model phosphatase–clay and phosphatase–metal (hydr)oxide complexes show a general loss in enzyme activity due to declines in the velocity of the reactions (Vmax) or a reduction in the affinity of the enzyme for its substrate (increased Km) (Gianfreda and Bollag, 1996; Huang et al., 1999; Quiquampoix and Mousain, 2005). Similar declines in velocity and affinity have also been noted with adsorption to model mucigel compounds (calcium–polygalacturonate) (Marzadori et al., 1998) and whole soil (PerezMateos et al., 1991; Gianfreda and Bollag, 1994; George et al., 2006). However, mixtures of soil components appear to have mitigating effects. For example, phosphatase associated with tannic acid had a reduced reaction velocity and substrate affinity, but when these complexes were in the presence of montmorillonite, velocity was unaffected and the affinity of the reaction was increased (Rao and Gianfreda, 2000). Different clay types also have differential effects on the kinetics of reactions. Montmorillonite reduced the velocity of phosphatase reactions, while kaolinite reduced substrate affinity (Dick and

Interactions Between Phytase and Soil Constituents

Tabatabai, 1987). Adsorption of phytase from A. ficuum (now known as A. niger) on gelatin particles had similar effects of declining velocity and affinity (Liu et al., 1999). Moreover, when phytases from two soil fungi (A. niger and P. lycii) were added to soil, both their velocity and substrate affinity declined (George et al., 2006). Of particular interest was that although velocity was not different between the two phytases in the soil environment, the affinity for inositol phosphate of the phytase in solution (P. lycii) was double that of the adsorbed (A. niger) enzyme (George et al., 2006). This suggests that greater proportions of phytase in solution increase the capacity for interaction with inositol phosphates. This was further demonstrated by the greater ability of P. lycii phytase, which is less adsorbed to soil surfaces than A. niger phytase, to mineralize inositol phosphates endogenous to a range of soils (George et al., 2006). Interestingly, it has also been demonstrated that the extracellular phytase from a range of soil fungi (Aspergillus spp., Emericella spp. and Penicillium spp.) is more effective at hydrolysing inositol phosphates compared with intracellular equivalents from the same organisms (Tarafdar et al., 2002), suggesting the existence of traits peculiar to the extracellular protein that allow more effective function in the soil environment. The observed effects of immobilization of phytases on their catalytic activity may depend on the following factors (Laidler and Sundaram, 1971; Ladd and Butler, 1975; Engasser and Horvath, 1976; Goldstein, 1976): 1. conformational change upon adsorption leading to loss of enzyme activity, reduced substrate specificity, or altered enthalpy/entropy; 2. changes caused by partitioning of the pH environment, substrate, products, activators and inhibitors between the enzyme’s microenvironment and the bulk soil solution; 3. steric hindrance by matrix shielding or occupation of active sites by inhibitors; and 4. effects of external and internal diffusion. Immobilization may impact the velocity of phytase reactions by causing conformational changes that render the enzyme denatured or reduce the rate of substrate turnover at the enzyme active site. Likewise, the entropy of immobilized phytase will be increased due to reduced flexibility, resulting in an enzyme less likely to achieve conformational requirements for hydrolysis. However, in

229

some cases immobilization may also reduce enthalpy in comparison to free phytase, resulting in a reduced energy requirement to reach the transition state (Kelleher et al., 2004). The balance between the effects of immobilization on enthalpy and entropy will, at least partially, determine the maximum velocity of the immobilized reaction. The affinity of an immobilized phytase for its substrate will only be the same as for the free enzyme when the supporting matrix is either uncharged or the ionic strength is great. When supports and substrates are similarly charged, as in the case of phytase immobilized on a clay surface, the inositol phosphate concentration will be reduced in the microenvironment of the enzyme, and the affinity, reduced. In contrast, if the support and substrate carry opposing charge, as is the case when phytase is immobilized on metal (hydr)oxides or some organic materials, the inositol phosphate is attracted to the microenvironment and the affinity of the reaction may increase (Crook et al., 1970; Ladd and Butler, 1975). Altered partitioning of inhibitors, activators and products of the catalytic reaction between the immobilized phytase and soil solution may also affect the catalytic efficiency of phytase. A further consequence of partitioning between immobilized phytase and soil solution will be an apparent shift in pH optima. This is explained by accumulation or dissipation of hydrogen ions in the enzyme’s microenvironment (Violante and Gianfreda, 2000), dependent on the charge of the supporting surface. These shifts in pH optima are therefore not due to changes in the properties of the immobilized enzyme per se, but are an artefact of the difference between measured pH and that in the microenvironment (Goldstein, 1976). Never the less, it is experimentally difficult to separate this potential pH surface effect from well-documented pH-dependent modifications of conformation or orientation of the enzyme on the solid surface (Baron et al., 1999; Quiquampoix 2000; Servagent-Noinville et al., 2000). Reduced activity of enzymes associated with high-molecular weight organic substances has, in most cases, been attributed to steric limitations to the penetration of substrates to the active site (Goldstein, 1976). It is likely that the affinity of phytase will vary depending on the soil particlesize fraction the enzyme is associated with (Marx

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et al., 2005). Although it has been suggested that different isozymes, with different affinities, are associated with various size fractions (Rojo et al., 1990), it is probable that enzymes associated with larger particle-size fractions have greater access to substrates than in smaller fractions. Therefore, steric hindrance is generally considered to be the main reason for reduced affinity with declining fraction size. Diffusion rates of both phytase and inositol phosphate are clearly important factors that restrict the affinity of their reaction in soil. Diffusion of free phytase will depend on its net charge and size as determined by the amino acid sequence and the degree of modification or substitution (Table 14.1). For example, greater glycosylation of fungal phytases compared to bacterial phytases may impact their relative mobility (Lei and Porres, 2003). The mobility of phytase itself may not be a major limitation to affinity towards an unlimited substrate. However, the availability of inositol phosphates in soil solution is likely to be limited (see Celi and Barberis, Chapter 13, this volume) and catalytic reactions will be reduced significantly by either the rate of diffusion of substrate to enzyme or vice versa (McLaren and Packer, 1970). Consequently, when diffusion rates are less than the rate of hydrolysis, a depression in phytase activity may be observed (Ladd and Butler, 1975; Goldstein, 1976; Ciurli et al., 1996). In soil, diffusion rates of inositol phosphate will depend on the soil type (sorption capacity for inositol phosphate) and also the tortuosity of path for diffusion, which in turn is dependent on water content. Importantly, the number and distribution of phosphate groups on the inositol ring will affect the rate of diffusion to, and within, enzyme supports (Engasser and Horvath, 1976) due to the reaction with anion exchange sites. This is likely to affect the specificity of immobilized phytase for the various inositol phosphates and stereoisomeric forms. The kinetics of an enzyme that is embedded in a porous matrix, such as an organic complex, will be further complicated by rates of internal diffusion, which will decrease with increasing depth into the matrix owing to progressive depletion of the substrate (Goldstein, 1976). Therefore, the distribution of enzymes in relation to the soil solid phase will affect the performance of the catalytic reaction (Nannipieri and Gianfreda, 1998). For example, phytases located on the external

surface of soil aggregates or organic matrices are likely to be less affected by diffusional limitations than those internal to the structure of the soil aggregate. This has been demonstrated by the fact that the affinity of adsorbed enzymes for substrate is much greater when associated with crushed soil than with intact aggregates (Brahms and McLaren, 1974). An important caveat is that when inhibition by diffusion and other factors, whether chemical or biological, is concurrent, the combined effect is an apparent amelioration of the initially observed inhibition. For example, if an inhibitor reduces the absolute activity of adsorbed enzyme and this reduces the difference between the rate of hydrolysis and rate of diffusion, diffusional inhibition of catalytic activity will appear to be increased (Goldstein, 1976). This also has the effect of apparently increasing the stability of an immobilized enzyme. Therefore, in a diffusionally limited system such as soil, phytase activity may appear to remain stable even though the protein has undergone considerable denaturation (Goldstein, 1976).

Microbial and proteolytic degradation Like all enzymes, phytases around plant roots will immediately encounter a repressive environment, being subject to potential microbial and proteolytic degradation upon exudation or loss from the cytoplasm (Tinker and Nye, 2000). Importantly, there are specific variations in the biochemical nature of phytases in relation to their susceptibility to proteolytic degradation, thought to be due to their vulnerability to conformational change (Simon and Igbasan, 2002). Protein degradation invariably occurs at exposed loops on the surface of the molecule and directed mutagenesis of A. fumigatus phytase has yielded variants that are considerably more resistant to proteolysis (Fig. 14.4; Wyss et al., 1999b). Phytases from different soil fungi also exhibit variation in their susceptibility to microbial degradation. For example, phytase from P. lycii was more susceptible to microbial degradation than that from A. niger (George et al., 2006). Glycosylation of phytases may also further affect their susceptibility to microbial and proteolytic degradation and is known to be highly variable (Wyss et al., 1999b), particularly when expressed in heterologous systems. As discussed earlier, protection from microbial and

Interactions Between Phytase and Soil Constituents

proteolytic degradation is afforded by immobilization on to the soil solid phase presumably as a result of steric hindrance towards degradative agents (Nannipieri et al., 1996; Rao and Gianfreda, 2000; George et al., 2006). Indeed, this stabilization of immobilized enzyme most likely contributes to the level of endogenous phytase activity that exists in soil. Moreover, it suggests that the soil environment exerts a selection pressure on the enzyme such that extended residence differentiates only the most robust or most protected proteins.

Inhibition and activation by ions and metabolites Most phytases require divalent cations for activity, which is thought to be due to their involvement in the active conformation of the phytases (Irving, 1980; Choi et al., 2001). Plant phytases tend to have a broader range of activators than those from microorganisms; for example, plant phytases are activated by magnesium, calcium or cobalt ions (Peers, 1953; Nagai and Funahashi, 1962; Gibbins and Norris, 1963; Chang, 1966;

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Fig. 14.4. Proteolytic susceptibility of wild-type and mutated Aspergillus fumigatus phytase. Shown is the activity of A. fumigatus wild-type phytase (●), A. fumigatus S126N phytase mutant (■), A. fumigatus R125L/S126N phytase mutant ( ) after incubation in the presence of proteolytic enzymes and A. fumigatus wild-type phytase (▲) after incubation with a proteolytic enzyme preparation that had been pre-treated at 90°C for 20 min. (From Wyss et al., 1999b. Reproduced with permission from the American Society for Microbiology.)

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Lolas and Markakis, 1977), whereas bacterial phytases are activated solely by calcium ions (Irving and Cosgrove, 1971a; Powar and Jagannathan, 1982; Shimuzu, 1992; Kerovuo et al., 1998; Kim et al., 1998a; Choi et al., 2001). In contrast, yeast (Saccharomyces spp.) phytase is activated by iron(II) and copper(II) ions (Nayini and Markakis, 1984, 1986), whereas other fungal phytases do not require any specific activation by a cation (Irving and Cosgrove, 1974). It is reasonable to assume that divalent cation activators such as these are abundant in soil and should not limit the activity of phytases, particularly in the pH range over which most phytases are active. In contrast, phytase activity can be inhibited by a wide range of metals, including Ag, Cd, Co, Cr, Cu, Fe, Hg, Mn, Ni, Pb, Sn, W and Zn (Peers, 1953; Yamada et al., 1968; Powar and Jagannathan, 1982; Nayini and Markakis, 1984; Svenson, 1986; Shimuzu, 1992; Hayes et al., 1999), with mixtures of metal ions having at least additive effects on acid phosphatase (Renella et al., 2003). Although the mode of inhibition by metal ions is not clear, it is suggested that they may compete with activators, cause precipitation of substrates, alter the active conformation of the enzyme or cause steric hindrance of substrate to the active site (Lolas and Markakis, 1977; Gianfreda and Bollag, 1996). In particular, metal ions that form insoluble sulphides are strong inhibitors (in the order Mn < Co < Cd < Cu < Hg < Ag), which suggests that inhibition occurs through interaction with sulphhydryl groups in the active site of the enzyme (Shaw, 1954; Juma and Tabatabai, 1977; Geiger et al., 1998b; Huang and Shindo, 2000b, 2001). Metal ions tend to reduce the velocity but increase the affinity of the reaction of extracellular enzymes, suggesting that they enhance the binding of inositol phosphate with the enzyme catalytic site (Huang and Shindo, 2000a,b, 2001). Many of these metals occur naturally in soil, albeit at concentrations that are unlikely to be inhibitory to phytases. However, concentrations of such metals could be considered inhibitory in some field sites acutely polluted by human activity. Inhibition by anions including phosphate (the reaction product), fluoride and arsenate is also evident and appears to be more potent against plant than microbial phytases (Nagai and Funahashi, 1962; Gibbins and Norris, 1963; Chang, 1966; Mandal et al., 1972; Chang and Schwimmer, 1977; Lolas and Markakis, 1977;

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Hayes et al., 1999). Due to subsequent pleiotropic effects, intracellular enzymes (most isolated plant phytases) are likely to be more tightly regulated by product accumulation than extracellular enzymes (most isolated microbial phytases), which generally show little or no regulation by phosphate (Nannipieri and Gianfreda, 1998). Inhibition by anions can also occur through precipitation of cationic activators, or by competitive inhibition with inositol phosphates for the enzyme active site (Irving and Cosgrove, 1971b). Phosphate is generally in low concentrations in solution and is depleted rapidly from the rhizosphere, so it is unlikely to inhibit phytase activity. Moreover, product inhibition is generally considered more effective against transcriptional regulation of phytases than direct inhibition of extracellular phytase activity in the soil (Bianchetti and Sartirana, 1967; Olander and Vitousek, 2000; Vats and Banerjee, 2004). Notwithstanding this, phosphate concentrations in soil solution may become inhibitory during waterlogging or following application of manure or mineral fertilizer. Other potential inhibitory anions such as arsenate and fluoride are not common in soil solution, whilst anions such as nitrate, sulphate and chloride do not appear to inhibit phosphatases (Juma and Tabatabai, 1977). In contrast, inhibition of microbial phytases by chelating agents such as citrate, oxalate, tartrate and ethylenediaminetetraacetate (EDTA) appear to be more acute than that of plant phytases (Irving and Cosgrove, 1971a; Nayini and Markakis, 1984; Shimuzu, 1992; Yoon et al., 1996; Kerovuo et al., 1998; Choi et al., 2001). Chelating agents could inhibit enzyme activity by binding with activators, although the effect may be mitigated through complexation with metals that would be inhibitory to phytases. The concentration of plant and microbial metabolites, such as citrate, oxalate and malate, is greatly enhanced within the rhizosphere (Tinker and Nye, 2000) and could therefore have a major effect on phytases within this zone.

Denaturation by soil environmental factors The pH environment in soil is likely to be more extreme and temporally variable than that of the cytoplasm. Despite this, the biological range of

phytases (Table 14.1) appears well able to cope with this environment, having optimal pH for catalytic reactions ranging from 2.2 to 8.6 (Irving, 1980; Nayini and Markakis, 1986). However, most discrete plant and microbial phytases have narrow single optima in the range of pH 3.5–7.5, and show significant declines in catalytic activity with small changes in pH on either side of this optimum (Peers, 1953; Nagai and Funahashi, 1962; Gibbins and Norris, 1963; Chang, 1966; Irving and Cosgrove, 1971a; Chang and Schwimmer, 1977; Lolas and Markakis, 1977; Basha, 1984; Nayini and Markakis, 1984; Greiner et al., 1993; Yoon et al., 1996; Kerovuo et al., 1998; Kim et al., 1998a; Hayes et al., 1999; Liu et al., 1999; Choi et al., 2001; Quan et al., 2004). Soil environments are unlikely to be conducive to optimal catalytic activity of phytases, as the pH is unlikely to be either optimal or remain stable at this optimal value. Notwithstanding this, phytase activity isolated directly from soil tends to have a broader range of pH optima (Svenson, 1986), and discrete phytases have been shown to have multiple and broad pH optima (Irving and Cosgrove, 1974; Greiner et al., 1993; Casey and Walsh, 2003; Brugger et al., 2004; Chadha et al., 2004; Dharmsthiti et al., 2005). Moreover, it is also now possible for phytases to be specifically engineered for broader pH optima (Mullaney et al., 2002; Tomschy et al., 2002). Although soil pH is unlikely to be optimal for phytase activity, it is also unlikely to lead to complete denaturation of the phytase protein, the structures of which are stable (activity was recoverable) against pH environments ranging from pH 1.2 to 11 (Yamada et al., 1968; Shimuzu, 1992). Phytases have generally been found to be temperature-stable with optimum activity in the range of 45–57ºC (Table 14.1). Beyond this, phytase activity tends to decline due to thermal denaturation with total denaturation occurring at ~80ºC (Irving, 1980). Importantly, some phytases (e.g. from A. fumigatus) are capable of re-forming their active conformation following exposure to high temperature (Wyss et al., 1998). Differences in glycosylation of phytases may also affect thermostability (Han et al., 1999), and specific modifications to the amino-acid sequence have been shown to increase tolerance to extreme temperature (Lehmann et al., 2000, 2002). Temperature is unlikely to denature phytases in soil under

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normal conditions, although exposed surface layers will approach or exceed denaturation temperatures in certain locations and certain times of the year. Total loss of phytase activity by thermal denaturation may also occur in soil during burning of above-ground vegetation (Saa et al., 1993; Staddon et al., 1998). This has implications for the recovery of biological cycling of inositol phosphate following natural forest fires or those used in agricultural and ecological management. Importantly, interaction with soil constituents can also make phytases less prone to thermal denaturation. For example, phytase isolated from mung bean was shown to be less sensitive to temperature denaturation when associated with inositol phosphates and divalent cations such as calcium (Mandal et al., 1972). Such interactions are assumed to alter the conformation of the enzyme, making it less susceptible to denaturation (Kim et al., 1998b; Choi et al., 2001). At typical soil temperatures standard thermodynamic principles will apply, such that enzyme kinetics will be temperature-dependent, with rates increasing up to the range of optimal temperatures mentioned above. Freezing may reduce the activity of extracellular phytases through denaturation (Pettit et al., 1977) and may affect the rates of enzyme reactions by changing the ionization of all reactants and the conformation of the protein (McLaren and Packer, 1970). The severity of freezing conditions is also important. Slow freezing generally leads to localized concentrations of reactants and an increase in the affinity of the system, whereas during rapid freezing the reactants remain homogenized and activity is severely retarded (McLaren and Packer, 1970). Water is essential for phytase activity, being the medium in which reactions occur. In general, enzyme activities tend to decline with drying (Gianfreda and Bollag, 1996). As soil dries, denaturation of extracellular enzymes occurs (Rao et al., 2003) due to unfolding of secondary structures. However, some enzymes regain their activity with rehydration, although self-association hinders the complete recovery of tertiary structures (Noinville et al., 2004). Although small changes in water potential may lead to modified protein structure and enzyme activity (Reyes et al., 2005), hydrolytic enzymes actually require very little water to be active. For example, extracellular urease requires 1.3 moles of water per mole of side chain polar groups to be active, suggesting that enzyme activi-

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ties, including phytase, may remain functional even in air-dried soil (McLaren and Packer, 1970). Interestingly, plant, microbial and fungal hydrophyllins (hydrophilic extracellular proteins) can reduce conformational changes to extracellular enzymes usually observed with reduced water potential, and thus avoid the loss of enzyme activity with drying (Reyes et al., 2005). A secondary effect of soil drying is an increase in ionic strength in soil solution. This may increase passive loss of phytases from microorganisms through osmotic stress and increase phytase activity in solution by reducing the adsorption of phytase through neutralization of electrostatic forces. Inhibitory or activatory effects of soil solution salts will also be concentrated by drying (Gianfreda and Bollag, 1996). In addition to dryness, excess water may impact phytase activity in soil. Waterlogging tends to limit extracellular enzyme activity (Freeman et al., 1996; Gianfreda and Bollag, 1996; Kang and Freeman, 1999; Chacon et al., 2005), through inhibition by metal ions such as iron and manganese in the reduced state, which are more soluble than their oxidized equivalents (Pulford and Tabatabai, 1988). As with metal ion toxicity, this effect appears to be mitigated by immobilization of extracellular enzymes on solid surfaces (Goel et al., 1998). Finally, phosphatases can be degraded by light, particularly short-wave radiation, such as ultraviolet-B (Espeland and Wetzel, 2001). This will be of little consequence to phytase in soil, except when enzymes are exposed to light either at the soil surface or following tillage (Nannipieri and Gianfreda, 1998). Photodegradation may be more important when phytases move from soil to aquatic environments. Despite the inherent longer exposure to light radiation following this transition, movement through the environment as organic–enzyme complexes will afford phytases some protection from photodegradation (Wetzel, 1992, 1993; Espeland and Wetzel, 2001) as will complexation with clay (Tietjen and Wetzel, 2003).

Concluding Remarks and Future Direction It is evident that the soil environment has a major effect on the ability of phytases to hydrolyse phosphorus from inositol phosphates. Phytases released

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to soil from plants and microorganisms, or those added to soil in manures from animals fed modified diets, will operate under suboptimal conditions. Major factors affecting phytase activity in soil include susceptibility to microbial and proteolytic degradation, variability of both pH and moisture content and, importantly, the immobilization and denaturation of phytase by the soil solid phase. Given these factors and that inositol phosphates react strongly in soil through precipitation and adsorption reactions, which can significantly affect their availability to phytases (see Celi and Barberis, Chapter 13, this volume), it is perhaps not surprising that inositol phosphates appear to resist mineralization and form a major constituent of the organic phosphorus in most soils. However, until we understand the role of the soil phytase activity that is retained in the long term and quantify the actual biological cycling of inositol phosphates in soil, it is difficult to speculate on the true importance of extracellular phytases in the soil environment. It remains to be proven whether such baseline phytase activity has an integral ecological role, such as warning microorganisms in their niche of temporal changes in the presence of inositol phosphates, as suggested for other phosphatases in Burn’s hypothesis, or whether these activities are simply fortuitously retained due to their protection against microbial degradation by adsorption to soil particles. However, the apparent interspecific variation in phytases and differences between functional groups (intracellular vs. extracellular) suggest that some advantage is gained by producing phytases with greater longevity in the soil environment, and that accidental retention of phytase activity would be a less-favoured conclusion. The challenge for future research is to better understand the efficiency of phytase–inositol phosphate interactions in soil. In particular, the following aspects require investigation: 1. the factors that control the availability of inositol phosphates for interaction with phytases; 2. the importance of the differing biochemical and physiological properties of phytases from various biological sources (e.g. bacteria, fungi, plants) and different classes of enzyme (e.g. histidine acid phosphatases vs. β-propeller phytase vs. purple acid phosphatases; see Mullaney and Ullah, Chapter 7, this volume) on the dephosphorylation of inositol phosphates in soil;

3. the role of phytases in the dephosphorylation of the range of inositol phosphates, including phosphorylated inositol stereoisomers, found in nature; and 4. the role of phytases in ecosystem function and their significance for the turnover of inositol phosphates as a component of the soil phosphorus cycle. Importantly, there is opportunity to exploit the natural variability in the biochemical characteristics of phytases or that which can be generated through protein engineering. Interspecific differences in the susceptibility of phytases to microbial degradation are evident, and the capacity to generate phytases that are less prone to proteolytic degradation is increasing. Similarly, phytases have been identified that are active over a range of soil pH and again it is possible that phytases with a broader range of pH optima can be engineered. Genetic variability in the susceptibility of phytases to immobilization by adsorption and subsequent degradation is also evident. Collectively, manipulation of these biochemical characteristics may make it possible to tailor specific phytases for optimal function in a range of soil environments and thus more effectively manage the interaction between phytases and inositol phosphates. The question remains, however, whether such changes would have any impact on the kinetics of phytase reactions in soil and thus the bioavailability of inositol phosphates. To address this it must be established whether soil–plant systems are already ‘optimized’ with respect to phytase activity and function at a ‘natural’ capacity, whereby the presence and/or accumulation of inositol phosphates over the long term is inevitable. Importantly, the ecological significance of phytase and inositol phosphates in soil–plant systems must be determined. We now have a range of experimental tools and analytical procedures that allow us to more thoroughly address some of these questions. For example, transgenic plants that express heterologous phytases can act not only as a delivery system for specific phytases to the rhizosphere, but also as bio-indicators to determine whether phytases with specific biochemical traits are effective at improving the bioavailability of inositol phosphates. This is important with regard to the utilization of inositol phosphates that are either

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endogenous to soil or added through animal manure and plant residues. At present, we are well poised to address key knowledge gaps in understanding critical parameters that control the turnover of inositol phosphates in soil. This will not only contribute to our

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understanding of the soil phosphorus cycle in both natural and agricultural ecosytems, but may also provide opportunity to improve phosphorus efficiency in agriculture, reducing the reliance on phosphorus fertilizer and any consequent environmental degradation.

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(1993) Changes in soil phosphorus and acid phosphatase activity immediately following forest fires. Soil Biology and Biochemistry 25, 1223–1230. Sarkar, J.M., Leonowicz, A. and Bollag, J.-M. (1989) Immobilisation of enzyme on clays and soils. Soil Biology and Biochemistry 21, 223–230. Servagent-Noinville, S., Revault, M., Quiquampoix, H. and Baron, M.H. (2000) Conformational changes of bovine serum albumin induced by adsorption on different clay surfaces: FTIR analysis. Journal of Colloid and Interface Science 221, 273–283. Shaw, W.H.R. (1954) The inhibition of urease by various metal ions. Journal of the American Chemical Society 76, 2160–2163. Shimuzu, M. (1992) Purification and characterisation of phytase from Bacillus subtilis (natto) N-77. Bioscience, Biotechnology, and Biochemistry 56, 1266–1269. Shimuzu, M. (1993) Purification and characterisation of phytase and phosphatase produced by Aspergillus oryzae K1. Bioscience, Biotechnology, and Biochemistry 57, 1364–1365. 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Plant Utilization of Inositol Phosphates

Alan E. Richardson1, Timothy S. George2, Iver Jakobsen3 and Richard J. Simpson1 1

CSIRO Plant Industry, PO Box 1600, Canberra, ACT 2601, Australia; Scottish Crop Research Institute, Invergowrie, Dundee DD2 5DA, United Kingdom; 3 Risø National Laboratory, Biosystems Department, Roskilde, DK 4000, Denmark

2

Inositol phosphates are a major component of the organic phosphorus in most soils, yet our understanding of the availability of inositol phosphates to plants is limited. Phosphorus deficiency is a major constraint to the growth of plants in many agricultural systems throughout the world and influences species composition in natural ecosystems (e.g. Wassen et al., 2005). This is despite the fact that soils generally contain a relatively large amount of total phosphorus. This total phosphorus includes organic and inorganic forms that are bound to soil particles by adsorption reactions or are present in mineral and precipitated complexes. Agricultural production systems are therefore reliant on the application of phosphorus-based fertilizers to meet the phosphorus requirement of plants. Such fertilizers are composed primarily of soluble inorganic phosphate processed from rock phosphates, or are derived from animal manures and other biological residues. However, much of the inorganic phosphate that is added to soil is rapidly ‘fixed’ (by adsorption and precipitation reactions) or is immobilized into organic phosphorus by soil microorganisms (Sanyal and De Datta, 1991; Oberson and Joner, 2005; Pierzynski et al., 2005), with the result that only a relatively small proportion of the phosphorus applied as fertilizer is taken up by plants. Consequently, there is the need to better understand how plants acquire phosphate from ‘endogenous’ forms of soil phos242

phorus, from applied sources of organic phosphorus, or from phosphorus that accumulates under different management systems. Accumulation of organic phosphorus and its utilization by plants are of particular interest, because organic phosphorus accounts for at least 50% and up to 80% of the total phosphorus in many soils (Harrison, 1987). Whilst much of the organic phosphorus in soil is associated with high-molecular weight fractions, a large part comprises phosphate monoesters. Of this, various stereoisomers of inositol penta- and hexakisphosphates are the major constituents and account for approximately 50% of the total organic phosphorus (Anderson, 1980; Turner et al., 2002b; see Turner, Chapter 12, this volume). Inositol phosphates are also the major storage compound for phosphorus in plant seeds, in which salts of myoinositol hexakisphosphate (phytate) account for ~70% of the total seed phosphorus (see Raboy, Chapter 8, this volume). Inositol phosphates (primarily as phytate) are thus a significant component of the dietary phosphorus intake of animals in intensive livestock industries. For monogastric animals in particular (i.e. swine and poultry) inositol phosphates in manures may therefore be important to the phosphorus cycle in soil–plant systems fertilized with manure (see Leytem and Maguire, Chapter 10, this volume). However, we have little understanding of the reactions of inositol phosphates in soil or their ‘biological

©CAB International 2007. Inositol Phosphates: Linking Agriculture and the Environment (eds B.L. Turner, A.E. Richardson and E.J. Mullaney)

Plant Utilization of Inositol Phosphates

availability’ to plants and microorganisms in soil environments. This chapter considers the importance of inositol phosphates as a potential source of phosphorus for plant nutrition. In particular, the activity of phytase in the rhizosphere and its contribution to the hydrolysis of inositol phosphates in soil is considered. Better knowledge of the role of inositol phosphates in the phosphorus nutrition of plants may improve our understanding of ecosystem function, and may also provide strategies to improve the efficiency of phosphorus-fertilizer use in different agricultural systems throughout the world.

Phosphorus Nutrition of Plants and Adjustment to Phosphorus Deficiency Organic forms of phosphorus are not directly available to plants, which take up phosphorus as soluble phosphate anions (HPO42− and H2PO4−). This occurs primarily across the plasma membrane of the root epidermis, through root hair cells or by mycorrhizae that are associated with roots. In most soils the concentration of phosphate in soil solution is low. Phosphorus deficiency occurs when the capacity for replenishment of phosphate or rates of diffusion are insufficient to meet plant requirements (Bieleski, 1973; Seeling and Zasoski, 1993). Plants have evolved a range of mechanisms that improve their capacity to acquire phosphate from the external environment and to maximize internal phosphorus utilization when deficient. These mechanisms are reviewed in detail elsewhere (e.g. Raghothama, 1999, 2005; Vance et al., 2003) and in summary include: 1. morphological changes to root structure such as rate of root growth, increased total and specific root length, the degree of root branching and the abundance and length of root hairs (Lynch, 2005; Hill et al., 2006), all of which allow plants to explore greater volumes of soil; 2. association with soil microorganisms, in particular mycorrhizal fungi and non-symbiotic microorganisms that can enhance either the availability or the uptake of phosphate from soil (Richardson, 2001; Jakobsen et al., 2005); and 3. biochemical processes that occur at the root–soil interface and within the rhizosphere. This includes induced expression of specific

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proteins for the transport of phosphate across the plasma membrane of root cells or those which facilitate its transfer from mycorrhizal fungi (Rausch and Bucher, 2002), modification to rhizosphere pH (Hinsinger, 2001), the release of root exudates (e.g. low-molecular weight organic anions) that improve phosphorus availability through increased solubilization of both inorganic and organic phosphorus pools (Hocking, 2001; Ryan et al., 2001) and the release of phosphatase enzymes (in particular acid phosphomonoesterases and -diesterases), which are required for the hydrolysis of organic phosphorus substrates (Richardson et al., 2005). Many plant species form symbiotic associations with mycorrhizal fungi, with the association generally being characterized by a mutualistic exchange of carbon from the plant in return for mineral nutrients from the soil, primarily phosphate (Smith and Read, 1997). Of particular note are the ectomycorrhizal fungi, which form associations predominantly with woody plants, and the arbuscular mycorrhizae, which associate with the majority of agricultural species. Characteristic of ectomycorrhizal infections is the formation of mycelial sheaths that envelop plant roots with hyphae that, although associated with the cell wall, are external to root cells. Arbuscular mycorrhizae have inter- and intracellular hyphae that penetrate the wall and plasma membrane of root cortical cells with the formation of haustoria-like arbuscules within plant cells. In both instances, it is well established that the fungal mycelia/ hyphae increase significantly the surface area of plant roots and provide greater contact with soil allowing enhanced uptake of phosphate (Jakobsen et al., 2005). There is little evidence to indicate that mycorrhizal fungi have access to pools of soil phosphorus other than those available to plants (Bolan, 1991; Joner et al., 2000), although it has been suggested that phosphatase activity in mycorrhizae may provide plants with increased access to soil organic phosphorus (Tarafdar and Marschner, 1994b; Feng et al., 2002).

Phosphatases and the utilization of soil organic phosphorus Organic phosphorus in soil and soil solution is not directly available to plants and must first be

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hydrolysed by phosphatase enzymes to release the phosphate anion. Dephosphorylation may occur either in the external root environment or, if substrates are soluble and able to diffuse through the root apoplast, within the cell wall space or at the root endodermis. Plants produce a range of extracellular acid phosphatases that are associated with either root cell walls (i.e. the root apoplasm) or released into the external environment as free enzymes (e.g. Dracup et al., 1984; Barrett-Lennard et al., 1993; Tadano et al., 1993; Li et al., 1997; Gilbert et al., 1999; Hayes et al., 1999; Zhang and McManus, 2000). Characterization of purple acid phosphatase genes (see Mullaney and Ullah, Chapter 7, this volume) from arabidopsis (Arabidopsis thaliana (L.) Heynh.) and white lupin (Lupinus albus L.) has confirmed the extracellular nature of these enzymes and their induced expression in response to phosphorus deficiency (Haran et al., 2000; Wasaki et al., 2000; Miller et al., 2001). Acid phosphatase activities have likewise been reported in isolated mycelium of mycorrhizal fungi. However, as with plant roots, cell-bound phosphatase activity generally dominates over extracellular forms (Straker and Mitchell, 1986; Kroehler et al., 1988; Ho, 1989; Antibus et al., 1992; McElhinney and Mitchell, 1993; Joner and Johansen, 2000, Olsson et al., 2002). Colonization of roots by both ectomycorrhizal and arbuscular mycorrhizal fungi has also been shown to increase the acid phosphatase activity of roots (Antibus et al., 1981; Dodd et al., 1987; MacFall et al., 1991; Ezawa and Yoshida, 1994; Fries et al., 1998), although its contribution to the extracellular component of root activity is unclear (Ezawa et al., 2005). Presence of extracellular acid phosphatase activity in hyphae associated with roots has been visualized histochemically for both arbuscular and ectomycorrhizae (Feng et al., 2002; Alvarez et al., 2005) and for intact plant roots (Dinkelaker and Marschner, 1992; Grierson and Comerford, 2000). Release of extracellular phosphatases from plant roots is also consistent with greater activity in the rhizosphere of soil-grown plants, particularly in response to phosphorus-deficient conditions (reviewed by Richardson et al., 2005). Greater phosphatase activity is generally accompanied by a depletion of soil organic phosphorus from the rhizosphere (Tarafdar and Jungk, 1987; Chen et al., 2002; George et al., 2002; Liu et al., 2004). Greater activity of acid phosphatases has simi-

larly been correlated with hyphal length of ectomycorrhizae associated with roots (Häussling and Marschner, 1989) and in some cases with the mycelial density of both arbuscular and ectomycorrhizae in root-free soil compartments (Tarafdar and Marschner, 1995; Feng et al., 2002; Liu et al., 2005), although this has not been observed in all cases ( Joner and Jakobsen, 1995; Joner et al., 1995). In the study by Liu et al. (2005) using radiata pine (Pinus radiata D. Don.), soil phosphatase activity was positively correlated with the length density of mycelium in root-free zones of soil and was associated with a significant depletion of soil organic phosphorus. However, the relative importance of phosphatases produced by plant roots, mycorrhizae or other free-living microorganisms in the rhizosphere, the activity and numbers of which are also substantially larger around roots ( Jakobsen et al., 2005), is not well understood. Whilst it is evident that phosphatases in the rhizosphere are effective for the depletion of organic phosphorus in various operationally defined pools (e.g. extractable in sodium bicarbonate, sodium hydroxide), there is a need to investigate the interaction of specific acid phosphatases and the utilization of defined organic phosphorus substrates (Richardson et al., 2005).

Implications for the utilization of inositol phosphates by plants in soil Despite the abundance of inositol phosphates in soil, their use by plants will depend on various factors that include: 1. The proximity to roots of inositol phosphates in soil. Roots (with or without mycorrhizae) must effectively explore soil to interact with substrate and to capture phosphate released by hydrolysis in competition with other reactions of phosphate in soil (e.g. immobilization by soil microorganisms or physical and chemical fixation). Roots and mycorrhizae are potentially well suited to exploit phosphorus in patchy environments such as organic layers, where hyphal proliferation and penetration into soil pores may be stimulated (Ravnskov et al., 1999; Gavito and Olsson, 2003; Hodge, 2004). 2. The solubility and mobility of inositol phosphates either within the soil solution or the root apoplasm. Soil solution contains a wide range of organic

Plant Utilization of Inositol Phosphates

phosphorus compounds (Wild and Oke, 1966; Martin, 1970), and phosphate monoesters, including inositol hexakisphosphates, have been identified as a component of soil leachate (Espinosa et al., 1999; Toor et al., 2003). However, their concentration in soil solution is likely to be small. Moreover, due to their high charge density, inositol phosphates are not expected to diffuse freely in soil solution and within plant cell walls. Inositol phosphates adsorb strongly to clays, metal oxides and organic matter with sorption capacities being equivalent to about 4 times that of phosphate anions on a per molecule basis (Anderson et al., 1974; Shang et al., 1992; Celi and Barberis, 2005; see Celi and Barberis, Chapter 13, this volume). Depending on pH, inositol phosphates also form sparingly soluble precipitates with a range of cations, with calcium and magnesium complexes being predominant under alkaline conditions and aluminium and iron complexes under acidic conditions (Jackman and Black, 1951). These reactions contribute to the stabilization of inositol phosphates in soil and are major factors that will affect its concentration in soil solution, mobility within the solution phase and susceptibility to enzyme hydrolysis (Tang et al., 2006). 3. The presence and activity of phytases of either plant or microbial origin, their location in or around plant roots and their capacity to effectively interact with substrate. At present, we have limited knowledge of the interaction of phytase with inositol phosphates in the soil–root environment and poor understanding of the rate-limiting steps for substrate dephosphorylation (see George et al., Chapter 14, this volume).

245

provided at concentrations that were up to 20fold higher than the phosphate concentration required for the maximum growth of each species. Similarly, wheat (Triticum aestivum L.) had limited ability to acquire phosphorus from sodium-phytate when compared to a range of other monoester and diester substrates. Ester-bonded phosphates other than phytate produced equivalent growth and phosphorus uptake as plants supplied with inorganic phosphate (Richardson et al., 2000). In both cases the inability to utilize phosphorus from phytate was considered to be associated with low levels of extracellular phytase that was released from roots into the external medium. This was confirmed by significantly improved plant growth and phosphorus uptake of subterranean clover seedlings when a purified phytase was added to the growth media (Hayes et al., 2000b). Inoculation of plants with microorganisms that possess phytase activity also improved the phosphorus nutrition of plants supplied with phytate. Phosphorus uptake by wheat, subterranean clover and a range of other plant species was significantly greater in the presence of an isolate of Pseudomonas sp. that was selected for extracellular phytase activity (Richardson and Hadobas, 1997; Richardson et al., 2000, 2001a). Growth promotion that is attributable to the phytase activity of bacteria has similarly been reported for plants inoculated with Bacillus amyloliquefaciens and a range of Burkholderia spp. (Idriss et al., 2002; Unno et al., 2005). Collectively, these studies indicate that plant roots do not possess an extracellular phytase activity that is effective for the utilization of phosphorus from inositol phosphates and that this inability can be complemented by the phytase activity of microorganisms.

Utilization of Inositol Phosphates by Plants Grown in Axenic Culture Phytase activity of plant roots Plants have limited capacity to access phosphorus from inositol phosphates relative to other organic substrates when grown under controlled conditions where availability of substrate is not expected to be limited (Hayes et al., 2000b; Richardson et al., 2001b). Using a number of grass and pasture legumes, Hayes et al. (2000b) showed that plants grown in sterile agar were unable to effectively obtain phosphorus from myo-inositol hexakisphosphate (supplied as sodium-phytate), even when

Phytase activity of roots has been measured for a range of plant species and has generally been shown to be absent or to constitute a small component only of the extracellular phosphatase activity of plant roots (Barrett-Lennard et al., 1993; Asmar, 1997; Bosse and Köck, 1998; Gilbert et al., 1999; Hayes et al., 1999; Richardson et al., 2000; Lung and Lim, 2006). In wheat seedlings, for example, phytase accounted for

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between 1% and 5% of the total phosphomonoesterase activity of root extracts (as determined using para-nitrophenyl phosphate as a model substrate), and although evident in whole root assays, was not detected as an extracellular activity in root exudates (Richardson et al., 2000). This was evident irrespective of plant phosphorus status and despite a twofold increase in the level of total phosphomonoesterase activity in exudates in response to phosphorus deficiency. Likewise, in tobacco roots (Nicotiana tabacum L.), phytase constituted 2.7% and 4.6% of the total activity in phosphorus-sufficient and -deficient plants, respectively, and although detectable in the external medium over 14 days, was not present as a significant root-released extracellular enzyme (Lung and Lim, 2006). Phytase activities were, however, present in both whole root extracts and root-associated cell wall fractions. A cell wall component of extracellular phytase has similarly been observed for subterranean clover without measurable release of enzyme activity (Hayes et al., 1999; George et al., 2004). White lupin also releases acid phosphatase from roots in response to phosphorus deficiency, but without significant release of phytase activity (Gilbert et al., 1999). The exception to these observations is the phytase activity in root exudates from a range of plant species reported by Li et al. (1997), where activities were substantial and equivalent to the total acid phosphatase. Lack of an exuded phytase activity from plant roots is consistent with observations of the in situ localization of phytase activity in maize (Zea mays L.) roots, where it was predominantly confined to the root endodermis (Hübel and Beck, 1996). Subsequently, two phytase genes (with similarity to histidine acid phosphatases) were cloned from maize and, while expression of these genes is consistent with their role in the mobilization of phytate in seeds, one of the genes was also expressed in the endodermis, pericycle and rhizodermis of mature roots (Maugenest et al., 1999). Analysis of this gene and protein, however, provided no direct evidence for its release as an extracellular enzyme. It was hypothesized that the role of the phytase was in the mobilization of endogenous phytate in plant roots, such as those deposited as phosphorus-rich globoids in pericycle and endodermis cells (Campbell et al., 1991; Van Steveninck et al., 1994; Hübel and Beck, 1996). More recently, Xiao et al. (2005) identified an extracellular phytase (MtPhy1)

from barrel medic (Medicago truncatula Gaertn.), which was shown to be secreted to the root cell wall. Whether this phytase is also released into the external soil environment remains to be determined, as does its effectiveness in allowing barrel medic to utilize inositol phosphates. The medic phytase is a purple acid phosphatase that has similarity to a phytase identified in soybean (Glycine max L. Merr.), which, from its pattern of expression in cotyledons, was considered to be involved in the mobilization of phytate during seed germination (Hegeman and Grabau, 2001). Nonetheless, the identification of an extracellular phytase in medic roots provides new opportunity for furthering our understanding of the functional significance of phytases in plant roots. Phytase genes from different sources have been expressed in plants to facilitate the understanding of their role in the utilization of inositol phosphates. Richardson et al. (2001a) showed that expression of the phyA gene from Aspergillus niger in arabidopsis improved the growth and phosphorus nutrition of plants supplied with sodiumphytate. This ability was associated with the release of phytase as an extracellular enzyme from the roots of the transgenic plants. In comparison, wild-type plants or control plants (which also expressed phytase but without a signal peptide for extracellular targeting of the enzyme) did not respond when supplied with phytate (Richardson et al., 2001a). Enhanced phosphorus nutrition of transgenic plants that release PhyA to the rhizosphere has since been demonstrated for tobacco and subterranean clover (George et al., 2004, 2005c). A 70-fold increase in the activity of exuded phytase resulted in significantly improved ability of the plants to acquire phosphorus from phytate (Fig. 15.1). The effectiveness of extracellular release of heterologous phytases in plants has similarly been demonstrated by expression of a consensus (fungal) phytase in transgenic potato (Solanum tuberosum L.; Zimmermann et al., 2003), the β-propeller phytase from B. subtilis (168phyA) in both tobacco and arabidopsis (Lung et al., 2005) and, more recently, the expression of the medic MtPhy1 phytase in arabidopsis (Xiao et al., 2005). In a number of cases, these phytases have been shown to be equally effective when expressed in plants either with constitutive promoters (i.e. the CaMV35S promoter) or with promoters derived from phosphate transport genes (e.g. the

Plant Utilization of Inositol Phosphates

myo-Inositol hexakisphosphate (sodium-phytate)

No P

247

Na2HPO4

Trifolium subterraneum

ex::phyA

Null segregant

ex::phyA

ex::phyA

Shoot dry weight (mg/plant)

28.1

40.7

51.8

47.9

Shoot phosphorus (µg P/shoot)

48.3

103.5

299.0

305.3

Exuded root phytase activitya (nKat/g root dry wt)

−

1.3

107.9

−

aActivity

for wild-type plants was 0.6 nKat/g root dry weight.

Fig. 15.1. Growth and phosphorus nutrition and activity of phytase exuded from the roots of transgenic Trifolium subterraneum. Shown are plants that release the Aspergillus niger phytase (ex::phyA) as an extracellular enzyme and its corresponding null segregant transgenic control line. Plants were grown for 28 days in sterile agar either without added phosphorus (No P) or with phosphorus supplied as sodiumphytate (myo-inositol hexakisphosphate) or disodium phosphate (Na2HPO4) at 0.8 mM (with respect to phosphate). (From George et al., 2004, in which experimental details are reported.)

promoter from the AtPht1,2 gene from arabidopsis), which direct gene expression predominantly to root hair cells and are induced under conditions of phosphorus deficiency (Mudge et al., 2003; Zimmermann et al., 2003; Xiao et al., 2005). These observations are significant, because they provide further evidence that plants do not have an innate ability to utilize phosphorus from inositol phosphates and, in soil environments, may be dependent on microbial-mediated mineralization.

Growth and Phosphorus Nutrition of Plants in Soil with Exogenous Substrate The ability of plants to utilize phosphorus from inositol phosphates has been of long-standing interest in plant nutrition (e.g. experiments by Rogers et al., 1940), and a number of key studies have

investigated their effectiveness compared to inorganic phosphorus for plants grown in sand or soil (Martin, 1973; Tarafdar and Claassen, 1988; Beck et al., 1989; Adams and Pate, 1992; Findenegg and Nelemans, 1993). These studies have generally shown that phosphate from myo-inositol hexakisphosphate is available to plants, but its availability is dependent on the level of substrate supply and the phosphorus-sorption characteristics of the growth medium. In quartz sand, with low phosphorus-sorption capacity, myo-inositol hexakisphosphate (supplied as sodium- or calcium-phytate) was equally available as inorganic phosphate to lupins (L. albus and L. angustifolius L.) when supplied ad libitum at 0.5 mM (Adams and Pate, 1992), and similarly was available to maize when applied at greater concentrations (10 mM and above), where the total amount of phosphorus supply was well in excess of plant requirements (Findenegg and Nelemans, 1993). However, when supplied at a lower rate (0.2 mM, and at a total phosphorus

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supply that was about 3 times that of the plants uptake requirements), calcium-phytate was ineffective relative to inorganic phosphate. Significantly, the phosphorus nutrition of maize plants grown under these conditions was improved when phytase was added to the sand medium, indicating that rate of hydrolysis, as well as availability of substrate, was important (Findenegg and Nelemans, 1993). Hydrolysis of phytate was also shown to occur in the sand at high rates of substrate supply, both without the addition of phytase and in the absence of plants, which indicated that microbial mineralization occurred. Inositol phosphates are less effective for the phosphorus nutrition of plants when added to soils. Whilst some studies have shown them to be equivalent to either other organic phosphorus substrates or to inorganic phosphate (Tarafdar and Claassen, 1988), it has generally been shown that plant acquisition of phosphorus from exogenously added phytate is poor, and essentially a function of the phosphate-sorption characteristics of the soil (Beck et al., 1989; Findenegg and Nelemans, 1993). This is shown in Fig. 15.2, where the growth of wild-type tobacco plants in two soils of differing phosphate-sorption capacity was restricted compared with that for equivalent rates of inorganic phosphate, with greater restriction occurring in the higher phosphorus-fixing soil (A. Richardson, 2006, unpublished data). This is consistent with observations that, compared with RNA, glycerophosphate and inorganic phosphate, phytate was poorly available to lupin plants in a low phosphorus soil (Adams and Pate, 1992) and that phytate was essentially not available to maize plants in three contrasting soils when supplied at different rates (Findenegg and Nelemans, 1993). The limited capacity of plants to access phosphorus from exogenously supplied phytate has also been demonstrated using radioactively labelled substrate. Martin and Cartwright (1971) showed no evidence for plant uptake of 32P from labelled myo-inositol hexakisphosphate by ryegrass (Lolium perenne L.) when supplied at a rate of either 20 or 200 mg P/kg soil in two high phosphorus-fixing soils. Substantial uptake occurred in low phosphorus-fixing sand, but only at the higher rate of phytate supply. Soil microorganisms may be important for plant access to inositol phosphates in soil irrespective of the fact that its reactivity (i.e., adsorption and precipitation) is a major factor determining

availability to plants. This is particularly so given that plants do not possess high intrinsic phytase activity, yet in many cases are able to utilize phosphorus from phytate when supplied at high rates (Fig. 15.2). Martin (1973) investigated the supply of labelled myo-inositol hexakisphosphate to wheat plants grown in soils that were either sterilized or re-inoculated with a mixed population of rhizosphere bacteria or specific isolates that possessed phytase activity. Uptake of 32P by the plants was essentially dependent on the rate of substrate supply, and no major differences were observed in the amount of 32P that was taken up by the plants in the various soil treatments. However, the radiolabel was incorporated into the soil microbial biomass in all soils and significant mineralization occurred through time. On the contrary, Hübel and Beck (1993) found no evidence for depletion of labelled phytate in the rhizosphere of maize. Improved phosphorus nutrition of a range of plant species supplied with phytate in a phosphorus-fixing sand-vermiculite medium has been observed after inoculation with soil microorganisms (Richardson et al., 2001b). Findenegg and Nelemans (1993) also showed that the availability of phosphorus from phytate to maize in three soils of differing phosphorus-sorption capacity was improved by the addition of phytase enzyme, albeit at rates and substrate concentrations that were up to tenfold greater than that required for plants grown in sand. More recently, using transgenic tobacco plants that release PhyA, George et al. (2005c) showed that plant phosphorus nutrition was increased by up to 52%, compared with a wild-type and a transgenic control, in two soils supplied with calcium-phytate. Absolute growth of these plants was, however, still significantly less than that for plants that received an equivalent amount of inorganic phosphate.

Significance of mycorrhizae for utilization of inositol phosphates The contribution of mycorrhizae to the phytase activity of plant roots and utilization of exogenously supplied substrate has similarly been investigated in controlled culture and soil-based experiments. Phytase activity has been detected in both arbuscular and ectomycorrhizal fungi (Theodorou, 1971; Bartlett and Lewis, 1973), and

Plant Utilization of Inositol Phosphates

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Calciumphytate

Nicotiana tabacum

Alfisol phosphate

No P

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250

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Calcium-phytate 200 Alfisol 100

0 No P

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Phosphate

Shoot dry weight (mg/plant)

Calcium-phytate 200 Spodosol

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0 No P

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Phosphorus applied (mg P/kg soil) Fig. 15.2. Growth of Nicotiana tabacum in an Alfisol and Spodosol fertilized with different rates of phosphorus (mg P/kg soil) as either inorganic phosphate or calcium-phytate, or grown without added phosphorus (No P). (From A. Richardson, 2006, unpublished data.) Shown is the growth of the plants in the Alfisol at 35 days and shoot dry weight in both soils after 48 days. The Alfisol is a high phosphorusfixing soil and the Spodosol is a low phosphorus-fixing sand (note the difference in applied rates of phosphorus). (Details of the soils are reported in George et al., 2005c.) Values are the mean of five replicates and the bars show one standard error.

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in pure culture phytate has been shown to be available to a range of mycorrhizal fungi (Antibus et al., 1992; Chen et al., 1999; Sawyer et al., 2003; Midgley et al., 2004). However, McElhinney and Mitchell (1993) showed that phytate was hydrolysed less effectively than inorganic pyrophosphate or β-glycerophosphate by ectomycorrhizal fungi. Utilization of phytate has also been observed for arbuscular mycorrhizae grown in monoxenic culture with roots (Koide and Kabir, 2000). However, like plant roots, it is evident that phytase from mycorrhizae does not appear to be a significant component of the extracellular activity and generally represents a small component only of the total phosphatase activity in mycelia of both arbuscular and ectomycorrhizal fungi (Mousain et al., 1988; Antibus et al., 1992; McElhinney and Mitchell, 1993; Firsching and Claassen, 1996; Colpaert et al., 1997). In the study by Colpaert et al. (1997) phytase activity was greater at the surface of ectomycorrhizal P. sylvestris roots than uncolonized roots. These activities were, however, less than total acid phosphatase and a similar trend was observed for field-collected roots from a number of different plant species (Antibus et al., 1997). Whilst the significance of these observations for the potential of mycorrhizae to directly utilize inositol phosphates in soil remains unclear, it is evident that their contribution to total phytase activity in roots and in soil is likely to be small, as has been reported for acid phosphatases (Joner and Johansen, 2000; reviewed by Joner et al., 2000). Despite this, their role may be important given that they provide an important interface between plant roots and the soil environment. A number of studies have shown that mycorrhizae can enhance the uptake of phosphorus by plants when supplied with various sources of organic matter and added organic phosphorus substrates (Jayachandran et al., 1992; Joner and Jakobsen, 1994, 1995; Perez-Moreno and Read, 2000; Tibbett and Sanders, 2002; Baxter and Dighton, 2005), although this does not necessarily imply their direct involvement in mineralization. For example, big bluestem grass (Andropogon geradii) plants colonized with arbuscular mycorrhizae used phytate (and a range of other organic phosphorus substrates) more effectively than nonmycorrhizal plants (Jayachandran et al., 1992). However, whether the phytate was hydrolysed by the mycorrhizae or by other soil microorganisms,

and the mycorrhizae simply assisted in its subsequent uptake, was not determined. In an attempt to separate these aspects, Tarafdar and Marschner (1995) used sterilized soil amended with phytate and inoculated with combinations of both arbuscular mycorrhizal fungi (Glomus mosseae) and a phytase secreting A. fumigatus. In this study, phosphorus nutrition of plants was greatest in soil inoculated with both microorganisms and this was accompanied with a reduction in soil organic phosphorus. Compartmentalized pots that only allow fungal access to soil amended with phytate have also been used to indicate a mycorrhizalinduced reduction in organic phosphorus in soil immediately adjacent to the root compartment, suggesting direct utilization of substrate by the fungus (Tarafdar and Marschner, 1994a,b; Feng et al., 2003). On the contrary, using a nonphosphorus-retentive perlite medium, Colpaert et al. (1997) showed that utilization of soluble phytate by P. silvestris was poor and that infection with two different strains of ectomycorrhizae provided no additional benefit. This occurred despite the provision of substrate at a relatively high concentration and observations that phytase activity was greater on the surface of mycorrhizal roots.

Plant Utilization of Endogenous Inositol Phosphates in Soil Few studies have specifically investigated the biological utilization of inositol phosphates that are endogenous to soil, and our understanding of their contribution to the phosphorus cycle and rate of turnover in soil is poor. To a large extent this is due to the lack of access to appropriate analytical technologies for their direct study in soil and because reactivity of inositol phosphates with soil constituents is a major factor that restricts their availability (e.g. Martin, 1973; Adams and Pate, 1992). Despite this, there is emerging evidence to suggest that inositol phosphates are biologically available, albeit to a limited extent. Early studies showed that inositol phosphates were mineralized in cultivated soils presumably as a result of the mixing and subsequent exposure of organic matter to soil microorganisms (Williams and Anderson, 1968). Subsequently, various microorganisms in soil that have potential to utilize phytate have been identified (Greaves and Webley,

Plant Utilization of Inositol Phosphates

Phytase-hydrolysable phosphorus in soil suspension (µg P/g soil)

1969; Cosgrove et al., 1970; Yoon et al., 1996; Richardson and Hadobas, 1997; Idriss et al., 2002; Unno et al., 2005; see Hill and Richardson, Chapter 5, this volume), with many of these being isolated from around plant roots. A number of recent studies have also demonstrated the presence of phytase-hydrolysable organic phosphorus (as determined using a range of enzyme preparations that may or may not be specific for hydrolysis of myo-inositol hexakisphosphate; see Richardson et al., 2005) in various soil fractions, including trace amounts in soil solution and water extracts, and larger pools within aqueous soil suspensions (Pant et al., 1994; Shand and Smith, 1997; Hayes et al., 2000a; Hens and Merckx, 2001; Turner et al., 2002a; Toor et al., 2003; Fig. 15.3). Whilst these studies do not provide direct evidence for microbial or plant utilization of inositol phosphates in soil, they do indicate its potential biological availability. A more direct approach to the utilization of inositol phosphates by plants has been the use of solution 31P nuclear magnetic resonance (NMR) spectroscopy to investigate the dynamics of organic phosphorus around roots. Using this approach, Chen et al. (2004) and George et al. (2006a) have shown that phosphate monoesters were depleted from a range of soils and that this

20

Initial soil

After plant growth

251

depletion was accompanied by greater acid phosphatase activity in the rhizosphere. In the study by Chen et al. (2004), depletion was most evident in soil re-planted with radiata pine (P. radiata) compared to ryegrass, and mineralization of both myo- and scyllo-inositol hexakisphosphates was demonstrated, with the decline of myo-inositol hexakisphosphate accounting for between 18% and 100% of the observed depletion of the phosphate monoesters around pine roots (Chen et al., 2004; Turner et al., 2005b). Comparable decreases were not observed in soil under the grass. The greater depletion of inositol phosphates in soils from radiata pine was considered to be due to the colonization of the pine roots by ectomycorrhizal fungi, as also suggested by others (Liu et al., 2004; Scott and Condron, 2004; Liu et al., 2005). In the study by Scott and Condron (2004) the ectomycorrhizal fungus had access to root-free compartments of soil where the mycelium decreased the total extractable soil organic phosphorus to a similar extent as did the fungus in combination with plant roots. This suggests that ectomycorrhizal fungi may play a dominant role, although the contribution of other soil microorganisms to the mineralization of soil organic phosphorus cannot be discounted. Decreases in organic phosphorus were also smaller in parallel Initial soil

After plant growth No plant

16

LSD = 2.3 (P < 0.05)

12

Wild-type Transgenic control

8

ex::phyA 4 0 Alfisol

Spodosol

Fig. 15.3. Phytase-hydrolysable organic phosphorus in soil suspensions of an Alfisol and Spodosol. (From T. George and A. Richardson, 2005, unpublished data.) Phytase-hydrolysable phosphorus was determined using a non-specific phytase on aqueous suspensions (1:10 w/v) of bulk soils either initially or after 28 days incubation in a glasshouse either without plants (No plant) or on soil collected from the rhizosphere (0–2 mm) of Trifolium subterranean plants that were wild-type transgenic control, or released Aspergillus niger phytase (ex::phyA) as an extracellular enzyme. (From George et al., 2005b.) Shown is the concentration of organic phosphorus that was deemed phytase-hydrolysable by incubation of soil samples (5 g) for 24 h with an excess of the non-specific phytase (Sigma Chemical Company, St Louis, Missouri, USA). Bars show one standard error of the mean (n = 4).

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treatments with lucerne (M. sativa L.) and ryegrass that were colonized by arbuscular mycorrhizal fungi (Scott and Condron, 2004). This is consistent with other observations that arbuscular mycorrhizae do not appear to play a significant role in the direct utilization of soil organic phosphorus above that of the capacity of plant roots alone (Joner and Jakobsen, 1995; Joner et al., 1995).

Growth and phosphorus nutrition of transgenic plants that release extracellular phytase Utilization of inositol phosphates in soil has also been investigated using transgenic plants that release A. niger phytase as an extracellular enzyme from roots. These plants are useful as they allow comparisons to be made with control plants that, apart from the release of the phytase, are genetically identical. Using phyA-expressing subterranean clover, George et al. (2006a) showed that phosphate monoesters were depleted in the rhizosphere of transgenic plants that expressed PhyA and that this depletion accounted for 15% of the total monoester phosphorus (of which ~25% could be identified as inositol phosphates) present in an Oxisol. Depletion of phosphate monoesters phosphates in this soil was accompanied by an increase in the alkali-extractable inorganic phosphate. When evaluated for growth in a range of different soils, however, the transgenic phyA-expressing subterranean clover did not show improved phosphorus nutrition over control lines, except when grown in a Vertisol that was high in total organic phosphorus and water-extractable organic phosphorus amenable to hydrolysis by a substrate-specific phytase (George et al., 2005b). In this Vertisol, phosphorus uptake by the transgenic plants was increased by between 21% and 31% over the controls and occurred irrespective of whether the soil was pasteurized to minimize the influence of soil microorganisms. Interestingly, the lack of phosphorus nutrition response of the transgenic plants in other soils (including the Oxisol) occurred despite the presence of significant amounts of inositol phosphates in each soil, as determined by solution 31P NMR spectroscopy and pools of organic phosphorus that were amenable to hydrolysis by phytase (George et al., 2005b). For example, in two of the soils (a low

phosphorus-fixing Spodosol and a high phosphorus-fixing Alfisol; Fig. 15.2) a significant depletion of phytase-hydrolysable phosphorus from aqueous soil suspensions occurred over 28 days in both transgenic and control plants and in soil that was incubated under the same conditions but without plants (Fig. 15.3; T. George and A. Richardson, 2005, unpublished data). Differences were evident in the net depletion of phytase-hydrolysable phosphorus between the two soils and greater depletion occurred in the rhizosphere of the transgenic line when grown in the Spodosol. These results indicate that either the measure of ‘phytase-hydrolysable’ phosphorus in these soils was not a good indicator of the availability of inositol phosphates to plants, or that microbially-mediated mineralization was a dominant process in these soils and that any benefit from plant-produced phytases was consequently minimal. This is consistent with the lack of growth response of transgenic subterranean clover and tobacco plants when grown in these two soils (George et al., 2004, 2005c) and highlights the need to better understand the importance of substrate availability and its interaction with microorganisms and plant roots in different soils. Irrespective of this, plant-exuded phytase can be significant for the phosphorus nutrition of plants in these soils, because growth of transgenic plants was enhanced over that of control plants when the soils were fertilized with either phytate or inorganic phosphate (George et al., 2004, 2005c). The response of plants to fertilization with inorganic phosphate (14–32% and 20–50% increase in shoot phosphorus content over wildtype controls for tobacco and subterranean clover, respectively) is of particular interest and suggests that phosphate addition may increase the availability of inositol phosphates to plants. This might occur through either displacement of adsorbed inositol phosphates, given that the counter-reaction (i.e. displacement of phosphate by inositol phosphates) has been observed (Anderson et al., 1974; Helal and Dressler, 1989), or through de novo synthesis by soil microorganisms. Using a 33P tracer, George et al. (2006a) demonstrated the rapid incorporation of labelled phosphate into alkali-extractable organic phosphorus, including microbial phosphorus and a pool that was amendable to hydrolysis by a non-specific phytase. Based on changes in specific activity of these pools it was further evident that transgenic plants that expressed phyA (compared

Plant Utilization of Inositol Phosphates

to control plants) preferentially depleted phosphorus from recently synthesized organic phosphorus. Collectively, these results suggest that microbial synthesis of inositol phosphates occurred in this soil and that plants that released the microbial-derived phytase had greater access to this source of phosphorus. However, more work to confirm this hypothesis is required and there is a need to further investigate the dynamics of inositol phosphates in soil, particularly in relation to the activity of soil microorganisms. Considerable variation in the biochemical properties of microbial phytases has been established (Wyss et al., 1999; Lassen et al., 2001; see Greiner, Chapter 6, this volume) and differential interaction of these various phytases in soil and their effectiveness in hydrolysing inositol phosphates has been demonstrated (George et al., 2005a, 2006b; see George et al., Chapter 14, this volume).

to be further investigation into such possibilities using a much wider range of plant species from different ecosystems, in addition to the few agricultural species that have been examined to date. Future research also needs to specifically address the following issues: ●

Conclusions and Future Research Directions Despite the abundance of inositol phosphates in soil, our understanding of their biological availability and contribution to the soil phosphorus cycle remains incomplete. It is evident that inositol phosphates are less available than phosphate diesters and other phosphate monoesters and are considerably less available than inorganic phosphate. Whilst this may largely be a consequence of the reactivity of inositol phosphates with soil constituents, it is also apparent that many plants do not have an innate capacity to directly utilize inositol phosphates in soil and appear to be dependent on microorganisms for their hydrolysis. This raises a number of interesting questions concerning the biological relevance of inositol phophate–phytase interactions in soil. In particular, it is paradoxical that plants have not evolved an ability to release phytase and directly utilize inositol phosphates, given the abundance of inositol phosphates in soils. Plants have evolved a wide range of other mechanisms to acquire soil phosphate when grown under conditions of low phosphorus availability. Either the lack of available substrate in soil solution has precluded selection pressure for such a trait to evolve in plants, or plants have evolved to rely on microorganisms for the hydrolysis of inositol phosphates in soil environments and within the rhizosphere (e.g. Unno et al., 2005). Obviously there needs

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A better understanding of how plants might utilize inositol phosphates in soil. This will require more detailed assessment of the role of phytases produced by mycorrhizae and other soil microorganisms and how they interact with plant roots in the rhizosphere. These studies will need to consider the interaction of the different phytases with the various stereoisomers of inositol hexakisphosphate that are found in soil (see Turner, Chapter 12, this volume). This should involve compartmentalized experimental systems that allow controlled access by either roots or fungal hyphae to soil, combined with a wider range of different plant species, including transgenic plants that have novel ability to release different phytases from their roots, along with manipulation of soil microbial populations. Such experiments also need to more closely resemble field situations (e.g. Schweiger and Jakobsen, 2000; Liu et al., 2005) and use analytical procedures such as solution 31P NMR spectroscopy that allow inositol phosphates to be appropriately identified and quantified separately from other constituents of the soil organic phosphorus. The development and application of analytical procedures that measure ‘biologically relevant’ pools of inositol phosphates in soil. At present we have little understanding of which components of inositol phosphates in soil are amenable to hydrolysis by phytases. Inositol phosphates dissolved in soil solution would be expected to be most available to plants and microorganisms, yet we have little information to support this. Whilst techniques such as NMR spectroscopy provide valuable insight into the total inositol phosphate content of soil, they currently provide little information on their biological availability. There is the need therefore to develop extraction or fractionation procedures that allow ‘meaningful’ pools of organic phosphorus to be identified (Turner et al., 2005a). Likewise, a better understanding of the biological relevance of phytase-hydrolysable pools, as determined by

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enzyme hydrolysis assays, is required. To date these too have proven to be of limited value in relation to the apparent availability of inositol phosphates to plants or microorganisms. Although transgenic plants that express phytase have shown significant phosphorus nutrition response in soils that have both large concentrations of total and waterextractable organic phosphorus that is amenable to hydrolysis by phytase (George et al., 2005b), these soil properties alone were not reliable indicators of the potential for plant response. Moreover, the understanding of how various chemical and physical attributes of soil interact with the availability of inositol phosphates is poor. Assessment of the fate of inositol phosphates in soil and factors that contribute to their synthesis and degradation within different components of soil biological systems. This will require quantitative analysis of inositol phosphate turnover in soil (e.g. using radioactive substrates) and capacity to differentiate ‘newly’ synthesized compounds (e.g. by microorganisms in the rhizosphere) from more stable forms that exhibit greater resistance to mineralization.

Recent observations showing differential interaction of various phytases with a range of metal ion–associated soluble and precipitated forms of phytate are significant (Tang et al., 2006), but such studies need to be extended to soil environments. The ability of microorganisms and plant roots to access inositol phosphates from these more recalcitrant forms, and to modify the chemical environment for its hydrolysis through the release of various exudates (e.g. organic acids; Hayes et al., 2000a; Tang et al., 2006) is important. Similarly, the fate of inositol phosphates that enter soil–plant systems through the application of animal manures and other organic phosphorus residues needs to be addressed. The contribution that inositol phosphates in soil make to the phosphorus nutrition of plants therefore remains somewhat uncertain and there is still much to learn concerning the biological interactions of inositol phosphates in terrestrial environments. This is important given the predominance of inositol phosphates in both agricultural and natural ecosystems.

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(2001) Function and mechanism of organic anion exudation from plant roots. Annual Review of Plant Physiology and Plant Molecular Biology 52, 527–560. Sanyal, S.K. and De Datta, S.K. (1991) Chemistry of phosphorus transformations in soil. Advances in Soil Science 16, 1–120. Sawyer, N.A., Chambers, S.M. and Cairney, J.W.G. (2003) Utilisation of inorganic and organic phosphorus sources by isolates of Amanita muscaria and Amanita species native to temperate eastern Australia. Australian Journal of Botany 51, 151–158. Schweiger, P.F. and Jakobsen, I. (2000) Laboratory and field methods for measurement of hyphal uptake of nutrients in soil. Plant and Soil 226, 237–244. Scott, J.T. and Condron, L.M. (2004) Short-term effects of radiate pine and selected pasture species on soil organic phosphorus mineralization. Plant and Soil 266, 153–163. Seeling, B. and Zasoski, R.J. (1993) Microbial effects in maintaining organic and inorganic solution phosphorus concentrations in a grassland topsoil. Plant and Soil 148, 277–284. Shand, C.A. and Smith, S. (1997) Enzymatic release of phosphate from model substrates and phosphorus compounds in soil solution from a peaty podzol. Biology and Fertility of Soils 24, 183–187.

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Shang, C., Stewart, J.W.B. and Huang, P.M. (1992) pH effect on kinetics of adsorption of organic and inorganic phosphates by short-range ordered aluminium and iron precipitates. Geoderma 53, 1–14. Smith, S.E. and Read, D. J. (1997) Mycorrhizal Symbiosis. Academic Press, San Diego, California. Straker, C.J. and Mitchell, D.T. (1986) The activity and characterization of acid phosphatases in endomycorrhizal fungi of the Ericaceae. New Phytologist 104, 243–256. Tadano, T., Ozawa, K., Sakai, H., Osaki, M. and Matsui, H. (1993) Secretion of acid phosphatase by the roots of crop plants under phosphorus-deficient conditions and some properties of the enzyme secreted by lupin roots. Plant and Soil 155/156, 95–98. Tang, J., Leung, A., Leung, C. and Lim, B.L. (2006) Hydrolysis of precipitated phytate by three distinct families of phytases. Soil Biology and Biochemistry 38, 1316–1324. Tarafdar, J.C. and Claassen, N. (1988) Organic phosphorus compounds as a phosphorus source for higher plants through the activity of phosphatases produced by plant roots and microorganisms. Biology and Fertility of Soils 5, 308–312. Tarafdar, J.C. and Jungk, A. (1987) Phosphatase activity in the rhizosphere and its relation to the depletion of soil organic phosphorus. Biology and Fertility of Soils 3, 199–204. Tarafdar, J.C. and Marschner, H. (1994a) Efficiency of VAM hyphae in utilisation of organic phosphorus by wheat plants. Soil Science and Plant Nutrition 40, 593–600. Tarafdar, J.C. and Marschner, H. (1994b) Phosphatase activity in the rhizosphere and hyphosphere of VA mycorrhizal wheat supplied with inorganic and organic phosphorus. Soil Biology and Biochemistry 26, 387–395. Tarafdar, J.C. and Marschner, H. (1995) Dual inoculation with Aspergillus fumigatus and Glomus mosseae enhances biomass production and nutrient uptake in wheat (Triticum aestivum L.) supplied with organic phosphorus as Naphytate. Plant and Soil 173, 97–102. Theodorou, C. (1971) The phytase activity of the mycorrhizal fungus Rhizopogon luteolus. Ecology 74, 1586–1593. Tibbett, M. and Sanders, F.E. (2002) Ectomycorrhizal symbiosis can enhance plant nutrition through improved access to discrete organic nutrient patches of high resource quality. Annals of Botany 89, 783–789. Toor, G.S., Condron, L.M., Di, H.J., Cameron, K.C. and Cade-Menun, B. J. (2003) Characterization of organic phosphorus in leachate from a grassland soil. Soil Biology and Biochemistry 35, 1317–1323. Turner, B.L., McKelvie, I.D. and Haygarth, P.M. (2002a) Characterization of water-extractable soil organic phosphorus by phosphatase hydrolysis. Soil Biology and Biochemistry 34, 27–35. Turner, B.L., Papházy, M. J., Haygarth, P.M. and McKelvie, I.D. (2002b) Inositol phosphates in the environment. Philosophical Transactions of the Royal Society, London, Series B 357, 449–469. Turner, B.L., Cade-Menun, B.J., Condron, L.M. and Newman, S. (2005a) Extraction of soil organic phosphorus. Talanta 66, 294–306. Turner, B.L., Mahieu, N., Condron, L.M. and Chen, C.R. (2005b) Quantification and bioavailability of scyllo-inositol hexakisphosphate in pasture soils. Soil Biology and Biochemistry 37, 2155–2158. Unno, Y., Okubo, K., Wasaki, J., Shinano, T. and Osaki, M. (2005) Plant growth promotion abilities and microscale bacterial dynamics in the rhizosphere of lupin analysed by phytate utilization ability. Environmental Microbiology 7, 396–404. Van Steveninck, R.F.M., Barbare, A., Fernando, D.R. and Van Steveninck, M.E. (1994) The binding of zinc in root cells of crop plants by phytic acid. Plant and Soil 155/156, 525–528. Vance, C.P., Ehde-Stone, C. and Allan, D.L. (2003) Phosphorus acquisition and use: critical adaptations by plants for securing a nonrenewable resource. New Phytologist 157, 423–447. Wasaki, J., Omura, M., Ando, M., Dateki, H., Shinano, T., Osaki, M., Ito, H., Matsui, H. and Tadano, T. (2000) Molecular cloning and root specific expression of secretory acid phosphatase from phosphate deficient lupin (Lupinus albus L.). Soil Science and Plant Nutrition 46, 427–437. Wassen, M. J., Olde Venterink, H., Lapshina, E.D. and Tanneberger, F. (2005) Endangered plants persist under phosphorus limitation. Nature 437, 547–550. Wild, A. and Oke, O.L. (1966) Organic phosphate compounds in calcium chloride extracts of soils: identification and availability to plants. Journal of Soil Science 17, 356–371. Williams, C.H. and Anderson, G. (1968) Inositol phosphates in some Australian soils. Australian Journal of Soil Research 6, 121–130. Wyss, M., Brugger, R., Kronenberger, A., Remy, R., Fimbel, R., Oesterhelt, G., Lehmann, M. and van Loon, A.P.G.M. (1999) Biochemical characterization of fungal phytases (myo-inositol hexakisphosphate phophohydrolases): catalytic properties. Applied and Environmental Microbiology 65, 367–373. Xiao, K., Harrison, M. J. and Wang, Z.Y. (2005) Transgenic expression of a novel M. truncatula phytase gene results in improved acquisition of organic phosphorus by Arabidopsis. Planta 222, 27–36.

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Yoon, S.J., Choi, Y.J., Min, H.K., Cho, K.K., Kim, J.W., Lee, S.C. and Jung, Y.H. (1996) Isolation and identification of phytase-producing bacterium, Enterobactor sp. 4, and enzymatic properties of phytase enzyme. Enzyme and Microbial Technology 18, 449 –454. Zhang, C. and McManus, M.T. (2000) Identification and characterization of two distinct acid phosphatases in cell walls of roots of white clover. Plant Physiology and Biochemistry 38, 259–270. Zimmermann, P., Zardi, G., Lehmann, M., Zeder, C., Amrhein, N., Frossard, E. and Bucher, M. (2003) Engineering the root–soil interface via targeted expression of a synthetic phytase gene in trichoblasts. Plant Biotechnology Journal 1, 353–360.

16

Inositol Phosphates in Aquatic Systems Ian D. McKelvie

Water Studies Centre and Chemistry Department, School of Chemistry, Monash University, Clayton, Victoria 3800, Australia

Although there is a considerable body of research on the abundance and behaviour of inositol phosphates in soils, much less is known regarding their behaviour in aquatic systems. This stems partly from the emphasis of aquatic research on the detection and measurement of molybdate reactive species in waters as a surrogate measure of bioavailable phosphorus. Inositol phosphates, like many other organic phosphate species, do not react with molybdate; consequently, they have been largely relegated to the fraction of phosphorus that is considered bio-unavailable, refractory and immobile. This view is overly simplistic and this chapter considers the known sources of inositol phosphates and likely transport paths and transformations in aquatic systems (Fig. 16.1; Turner et al., 2002b). The dominant inositol phosphate in soils and sediments appears to be myo-inositol hexakisphosphate, with the lower-order inositol phosphates occurring only as intermediates in either hydrolytic or biosynthetic sequences. This chapter will therefore focus on the behaviour of myo-inositol hexakisphosphate in aquatic systems. Suggested mechanisms for the release and transport of both inorganic and organic phosphorus from sediments are reviewed, and some speculative interpretation of the release, hydrolysis and bioavailability of inositol phosphates is offered.

Sources of Inositol Phosphates in the Aquatic Environment Inositol phosphates in aquatic systems are thought to originate from external, terrestrial sources such as soil particles and plant matter, or from internal sources such as algae and macrophytes. In general, plants contain only myo-inositol hexakisphosphate; so it seems likely that the scyllo-, D-chiroand neo-inositol phosphates are of microbial origin (Cosgrove, 1980), formed by epimerization from either myo-inositol or its hexakisphosphate (L’Annunziata, 1975). The phosphorylated inositol stereoisomers are discussed in detail elsewhere in this volume (see L’Annunziata, Chapter 4, and Turner, Chapter 12). Weimer and Armstrong (1979) studied the composition of inositol phosphates in several species of aquatic plants, algae and sediments of two fresh water lakes in Wisconsin, USA. For aquatic macrophytes and angiosperms, they found that lower-order esters of myo-inositol phosphate (i.e. tetrakisphosphate to monophosphate) were present in greater amounts than the hexa- and pentakisphosphate esters. They also noted that the ratio of higher-order to lowerorder inositol phosphates for catchment soils was greater than that found in the sediments, and suggested that this was due either to hydrolysis of soil inositol phosphates during transport into

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Fig. 16.1. Suggested sources, pathways and transformations of inositol hexakisphosphate (IP6) in aquatic ecosystems. (Modified from Turner et al., 2002b.)

the lake or to a much larger contribution of inositol phosphates from plant sources than expected. They concluded, however, that inositol phosphates in these lakes were derived approximately equally from algal primary production and catchment inputs, rather than from aquatic plants. These early findings were subsequently questioned, because the extraction conditions used (pH <1, 110°C, 6 h) may have dephosphorylated higher inositol phosphates (Potman and Lijklema, 1983). However, there are other reports of in vivo biosynthesis of inositol phosphates by aquatic plants, so this may be an important autochthonous source. For example, the aquatic angiosperm Wolffiella floridana was found to convert a high proportion of radiolabelled myo-inositol to inositol hexakisphosphate and lower inositol phosphates rather than to cell wall polysaccharides (Roberts and Loewus, 1968). Significant amounts of myoinositol hexakisphosphate were also found in the duckweed Spirodela polyrhiza during a developmental stage (Brearley and Hanke, 1996a), and the sequence of phosphorylation in this process was subsequently described (Brearley and Hanke, 1996b). Phosphorylation of myo-inositol to produce myo-inositol hexakisphosphate has also been reported to occur in the lesser duckweed Lemna minor (Inhulsen and Niemeyer, 1978) and the slime mould Dictostelium (Stephens and Irvine, 1990). Inositol phospholipids and lower-order inositol phosphates have also been implicated in phytoplankton metabolism and structure (Oku

and Kamatani, 1995). However, in a study of the coastal sediments in Tokyo Bay, Japan, Suzumura and Kamatani (1995b) found very little inositol phosphate in zooplankton and algae and reported that the major source of inositol phosphates was soils from surrounding catchments. This conclusion was based on the observation that the order of abundance of the myo-, scyllo-, and chiro-inositol phosphates were the same in sediments as in terrestrial and river suspended particulate samples. Hence, if biosynthesis were a major source of inositol phosphates in these sediments, it would be likely that varying ratios of inositol phosphate stereoisomers would be detected. The input of inositol phosphates from the manures of monogastric animals (poultry, swine) into aquatic systems is also a potentially large source of inositol phosphate, especially from catchments containing intensive production (e.g. feedlot farming). The manures of monogastric agricultural animals can contain high concentrations of myo-inositol hexakisphosphate because such animals have low levels of intestinal phytase (e.g. Maguire et al., 2004), and for this reason phytase is increasingly used as a feed additive to improve utilization of inositol phosphates and to reduce the requirement for inorganic phosphate supplements in animal diets (Valaja et al., 1998; Bedford, 2000; Juanpere et al., 2004; see Lei and Porres, Chapter 9, this volume). Improvement in dietary phosphorus availability through the use of phytase also appears to be beneficial by reduc-

Inositol Phosphates in Aquatic Systems

ing the amount of inorganic phosphorus excreted and added to catchments from manures (Penn et al., 2004; see Leytem and Maguire, Chapter 10, this volume). An extension of this approach has been the development of transgenic pigs that produce phytase in their saliva (Golovan et al., 2001). This enables them to digest phytate in the diet and is claimed to reduce faecal phosphorus output by up to 75%. Manure from poultry and swine not fed phytase-amended diets is likely to contain high concentrations of myo-inositol hexakisphosphate (although see Leytem et al., 2004), but it is poorly soluble and would likely be strongly bound to soils (Turner and Leytem, 2004). However, despite its low solubility, transport into streams in either colloidal or particulate forms might reasonably be expected under overland flow conditions (Turner, 2005).

Physicochemistry of Inositol Phosphates in Aquatic Systems myo-Inositol hexakisphosphate is resistant to chemical hydrolysis, especially under alkaline conditions. The maximum rate of hydrolysis occurs near pH 4 (Cosgrove, 1980) and decreases to a minimum at pH 0–1. Less than 50% hydrolysis was achieved at pH < 0 and 100°C for 6 h, (Cosgrove, 1980), whereas at 50°C hydrolysis was <20% after 35 days (Potman and Lijklema, 1983). This behaviour has important implications for both the extraction of inositol phosphates from sediments and soils, and the analysis of the organic phosphorus component of sediments, pore waters and the overlying water column. Extraction with strong acids and higher temperatures for extended periods introduces the risk of myo-inositol hexakisphosphate hydrolysis (Potman and Lijklema, 1983, Turner et al., 2005a), which has been cited as a possible explanation for the high concentrations of lower-order inositol phosphates reported in lake sediments by Weimer and Armstrong (1979). On the other hand, digestions for total and total filterable phosphorus determination using nitric acid alone or nitric and sulphuric acids may give incomplete recovery of myo-inositol hexakisphosphate, especially if typical digestion times of less than 2 h are used. For complete conversion, inclusion of an oxidizing agent such as peroxydisulphate or hydrogen peroxide in the digestion reagent is recommended (Benson et al.,

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1996). This also highlights the need for adequate validation of the extraction, digestion and detection stages in the determination of inositol phosphates using appropriate model organic phosphorus compounds (Kérouel and Aminot, 1996). Abiotic reactions of inositol phosphates with charged surfaces and polyvalent cations are discussed in detail elsewhere in this volume (see Celi and Barberis, Chapter 13) and only a brief overview of aspects relevant to aquatic systems is provided here. myo-Inositol hexakisphosphate readily complexes multivalent metal ions, and the order of relative complex stabilities is thought to be: Cu2+> Zn2+ > Ni2+ > Co2+ > Mn2+ > Fe3+ > Ca2+ (Cosgrove, 1980). It also forms stable surface complexes with minerals such as goethite and Fe(OOH) (De Groot and Golterman, 1993), and with clay minerals such as illite and kaolinite. Adsorption may also disperse clay minerals by alteration of surface charge (Celi et al., 1999). The sorption of myo-inositol hexakisphosphate on goethite was observed to depend both on the nature of dissolved cations present and the pH. In the presence of potassium ions, adsorption was pronounced at low pH, but as pH increased the adsorption decreased in response to the increasing charge density of myoinositol hexakisphosphate (−8 at pH 5.5). Adsorption on to goethite (Fe(OOH)) also displaced phosphate that was already adsorbed, and prevented further adsorption of phosphate. As was the case for the clay minerals, myo-inositol hexakisphosphate adsorption had a pronounced effect on the surface charge, resulting in dispersion of goethite particles. When calcium ions are present, however, adsorption continues to occur even at higher pH, apparently due to precipitation of calcium-phytate complexes. In this case, the effect of myo-inositol hexakisphosphate adsorption is insufficient to cause particle dispersion, and aggregation of goethite occurs in the presence of Ca2+ (Celi et al., 2001; see Celi and Barberis, Chapter 13, this volume). De Groot and Golterman (1993) also studied the effect of iron(III) reduction on myo-inositol hexakisphosphate adsorbed on goethite and reported that rather than being released in a solubilized form, myo-inositol hexakisphosphate remained bound as insoluble Fe4-phytate. Calcite is also reported to have a high capacity for retention of myo-inositol hexakisphosphate; this appears to involve a combination of adsorption and the complexation of calcium ions, accompanied by the dis-

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solution of calcite and the precipitation of Ca3-phytate (Celi et al., 2000). These adsorption and aggregation/dispersion effects have important implications for the accumulation of inositol phosphates in soils and colloidal transport in waters. The observed physicochemical behaviour of myo-inositol hexakisphosphate suggests that in marine and fresh waters it should be stable, complexed and/or precipitated by major ions such as calcium, magnesium, iron and manganese, or bound to the surfaces of suspended particles and thus immobilized in the sediments (De Groot and Golterman, 1993). In addition, inositol hexakisphosphate is thought to be associated with high-molecular mass humic acids (Hong and Yamane, 1980; Golterman et al., 1998).

Occurrence of Inositol Phosphates in Aquatic Systems Waters A commonly held view is that inositol hexakisphosphate exists in aquatic systems as an insoluble, refractory, immobile and biologically unavailable phosphorus species. Inositol hexakisphosphate is not hydrolysed by exocellular alkaline phosphatase or phosphodiesterase (Turner et al., 2002a), and is therefore regarded as biologically unavailable to phytoplankton. Thus it might be expected that the concentration of inositol phosphate in solution in both pore and overlying waters would be very low, or even undetectable. However, Eisenreich and Armstrong (1977) used a combination of alkaline bromination and gel filtration to perform selective detection of inositol phosphates, and reported that there was between 3 and 15 µg P/l of this organic phosphorus species in the waters of Lake Mendota, Wisconsin, USA, which represented between 20% and 30% of the total filterable phosphorus present (Table 16.1). Similarly, substantial concentrations of high-molecular mass phosphorus, with similar elution times as inositol hexakisphosphate on a gel filtration column, have been detected in the filterable fraction of waters and sediment pore waters (Fig. 16.2) from south-east Australia (McKelvie et al., 1993; McKelvie, 2005). Concentrations of this fraction in a range of waters were between undetectable and 52 µg P/l, and in

some cases comprised most of the filterable phosphorus present. Although the resolution of the gel filtration separation was limited, the dominant highmolecular mass phosphorus peak always coincided with that for an authentic standard of myo-inositol hexakisphosphate; so it is reasonable to assume that this peak consisted predominantly of higher-order inositol phosphates. Others have used hydrolytic techniques based on the enzyme phytase to hydrolyse organic phosphorus in waters and sediment extracts (Cooper et al., 1991). For example, it was shown that up to 50% of the organic phosphorus in the waters of two small mesotrophic and hypereutrophic lakes was amenable to hydrolysis by phytase, and on the basis of gel filtration separations, that this organic phosphorus consisted either of soluble inositol phosphates or inositol phosphates associated with higher-molecular mass proteins, lipids or fulvic acid (Herbes et al., 1975). The same enzyme, 3-phytase, was used in an immobilized form in an automated flow injection system to determine the concentration of phytase-hydrolysable phosphorus in waters (McKelvie et al., 1995). Concentrations in a range of waters in south-east Australia were between 1 and 75 µg P/l, with the higher values being associated with estuarine waters. The determination of inositol phosphates by these approaches is, however, more inferential than definitive. The specificity of commercial 3-phytase preparations is poor; it catalyses hydrolysis of a wide range of phosphomonoesters and even some diesters (McKelvie et al., 1995). Consequently, concentrations reported as myoinositol hexakisphosphate by this approach will overestimate the true concentration. Similarly, the poor selectivity of low-pressure gel filtration separations means that, at best, separated peaks, such as those shown in Fig. 16.2, represent a size or mass range rather than a single species such as myo-inositol hexakisphosphate. The use of analytical techniques with high selectivity for inositol phosphates, such as that described by Clarkin et al. (1992) or Suzumura and Kamatani (1993), is preferable. For example, Espinosa et al. (1999), using high-performance ion exchange chromatography, obtained highly resolved separations and showed that myo-inositol hexakisphosphate constituted nearly one-third of the identifiable organic phosphorus compounds present in leachate from a temperate grassland soil.

Table 16.1. Indicative concentrations of inositol hexakisphosphate in water, using a variety of estimation methods. Inositol hexakisphosphate

(µg P/l)

Frain’s Lake, USA (highly eutrophic) Third Sister Lake, USA (eutrophic) Lake Mendota, Wisconsin, USA River (4) and estuarine (5) waters, rural Victoria, south-east Australia Urban river (2) and lake waters (2), Melbourne, Australia Yarra River sediment, south-east Australia

3.5–12.4

Overlying water Pore water

Fraction of total filterable P (%)

Method of detection

Reference

12–47a

Phytase hydrolysis, photometric detection of reactive phosphate

Herbes et al. (1975)

3–15

20–30

1–75 (mean 25)

15

Alkaline bromination, followed by gel filtration on Sephadex G25 Phytase hydrolysis, flow injection detection of reactive phosphate

Eisenreich and Armstrong (1977) McKelvie et al. (1995)

21–52 (mean 36)

83

Gel filtration on Sephadex G25, flow injection detection of reactive phosphate after online photooxidation Gel filtration on Sephadex G25, flow injection detection of reactive P after online photooxidation

McKelvie et al. (1993)

4.9–10.0

27 29–281 (mean 85)

McKelvie (2005)b

Inositol Phosphates in Aquatic Systems

Sample origin and type

66 85

a

Proportion of the total organic phosphorus. See Fig. 16.2 for further details.

b

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I.D. McKelvie

(a)

0−1 cm core segment

3−4 cm core segment

60

16 12

40

8 20 4 0

0

2

4

6

8

0

0

2

Time (min)

4

6

8

Time (min)

(b) 300

Phosphorus concentration (µg P/I)

LMMP HMMP 250

200

150

100

50

0 Overlying water

0−1

1−2

2−3

3−4

4−5

5−7.5

Pore water depth (cm) Fig. 16.2. (a) Low-pressure, gel filtration separations of high- and low-molecular mass organic phosphorus (HMMP and LMMP, respectively). myo-Inositol hexakisphosphate and phosphate eluted at ~2.8 and 5.5 min, respectively. (b) Concentrations of phosphorus found in pore waters from Yarra River (Fairfield) sediment. (Redrawn from McKelvie, 2005.)

Sediments Given the apparent refractory and insoluble nature of metal-complexed myo-inositol hexakisphosphate, it might be expected that sediments would contain substantial amounts of phosphorus in this form. This is supported by reports that it may comprise as much as 80% of the total organic phosphorus (Weimer and Armstrong, 1977). However, the amount detected will be

highly dependent on the respective efficiency and the selectivity of the extraction and detection methods used (Table 16.2). Historically, soils and sediments were extracted with dilute acid to remove calciumbound phosphate, followed by strongly alkaline media such as hot 3 M NaOH to recover organic phosphorus, with final precipitation of inositol phosphates as barium salts (reviewed in Turner et al., 2002b). Further isolation of

Table 16.2. Reported concentrations of myo-inositol hexakisphosphate in sediments. Inositol hexakisphosphate

mg P/kg dry wt

Fraction of total P (%)

Sediments from ten lakes, Wisconsin, USA

–

51–80a

Sediments from Lake Mendota, Wisconsin, USA

49 38

12.1 9.3

51

12.7

Sediments from Tokyo Bay, Japan

1.9–6.2 (mean 3.7)

0.39 (mean)

Riverine and estuarine suspended solids from Tokyo Bay, Japan Sediments from Tokyo Bay, Japan

2.2–20.4 (mean 8.8)

0.75 (mean)

0.3–3.1 (mean 1.9)

0.22 (mean)

Sediments from Lake Wellington, Australia

2.5–14 (mean 6.3)

53 (mean)

Marsh and lake sediments from Camargue, France

24–149 (mean 86)

17.6a (mean)

Extraction and analytical methodology

Reference

0.3 M NaOH, 25ºC, 16 h, and 0.3 M NaOH, 100ºC, 8 h; ion exchange chromatography 3 M NaOH, 100ºC 1 M NaOH, 60ºC, hypobromite oxidation, barium precipitation 3 M NaOH, 100ºC, hypobromite oxidation, barium precipitation; ion exchange chromatography Hypobromite oxidation; ion exchange chromatography; gas chromatography Hypobromite oxidation; ion exchange chromatography; gas chromatography Hypobromite oxidation; ion exchange chromatography; gas chromatography 25 mM Na-tetraborate, pH 9.2, measured as high-molecular mass phosphorus using gel filtration with flow injection analysis 0.5 M HCl, 30 min, 2 M NaOH, 90ºC, 30 min, H2SO4, pH < 2, phytase

Sommers et al. (1972)

Weimer and Armstrong (1977)

Suzumura and Kamatani (1993) Suzumura and Kamatani (1995b) Suzumura and Kamatani (1995b) McKelvie et al. (1993)

Inositol Phosphates in Aquatic Systems

Sample origin and type

De Groot and Golterman (1993)

a

Proportion of the total organic phosphorus.

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myo-inositol hexakisphosphate can be achieved by the use of ion exchange chromatography (McKercher and Anderson, 1968), although this approach lacks selectivity because other organic phosphorus species may also be present. To overcome this, Irving and Cosgrove (1981) used bromine to oxidize all soil organic matter and organic phosphorus other than the higher-order inositol phosphates. However, it has been shown subsequently that while this procedure gives good recovery for myo-inositol hexakisphosphate (Weimer and Armstrong, 1977), it does not oxidize DNA (Nanny and Minear, 1994).

The efficiency of myo-inositol hexakisphosphate extraction of soils was shown to improve if higher temperatures were employed in the sodium hydroxide extraction step and longer time was allowed for the hypobromite reaction (Hong and Yamane, 1980). To avoid possible degradation of the sample by high temperature extraction a milder procedure was recommended for sediments (De Groot and Golterman, 1993). This involved extraction with complexing reagents such as ethylenediaminetetraacetate (EDTA) and dithionite, followed by extraction of acid-soluble and residual organic phosphorus (containing inositol phosphates) with 2 M NaOH (Fig. 16.3).

Pellet 0 (Total P)

Ca-NTA 0.02 M Dithionite Extraction of inorganic-P

pH = 7.8−8.0

Na-EDTA 0.05 M pH = ~8.0

Fe(OOH)~P Amorphous Fe(OOH)

CaCO3~P CaCO3

Pellet I (Organic P)

0.5 M H+ (HCI or H2SO4) 30 min

ASOP (Acid-soluble organic P)

Pellet II (ROP = residual organic phosphate)

2.0 M NaOH 90°C NaOHextr~P 30 min H2SO4 Pellet III (Rest~P)

pH < 2.0 Phytase Fulvic acid~P

Pellet IV (Humic acid~P)

Phytate~P (Inositol phosphate)

Fig. 16.3. Scheme for phosphorus fractionation in sediments, including the determination of inositol hexakisphosphate in the residual organic phosphate component. (Reproduced from De Groot and Golterman, 1993 with permission from Springer-Verlag.)

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Inferential evidence of the breakdown of inositol phosphate in sediments was also provided by a study that isolated and quantified inositols in marine sediments (White and Miller, 1976). While the proportions of unphosphorylated myo-, chiro- and scyllo-inositols were similar with depth, the total inositol concentration decreased. This was attributed to the effects of bacterial action and leaching on inositol phosphates. Further studies of Tokyo Bay sediments (Suzumura and Kamatani, 1995a) showed that under anaerobic conditions inositol hexakisphosphate was almost completely mineralized within 40 days, while under aerobic conditions it took 60 days for ~50% decomposition to occur. Decomposition under anaerobic conditions was ascribed to bacterial hydrolysis, and was considered to be more pronounced in the marine environments compared with fresh water environments. Depth profiles of myo-inositol hexakisphosphate in Yarra River pore waters (measured as high-molecular mass phosphorus; Fig. 16.2) showed a similar trend to that in Fig. 16.4 (McKelvie, 2005), although the presence of measurable myo-inositol hexakisphosphate in deeper sediments suggested that the rate of decomposition or removal is not as fast as that in marine systems. In a recent paper, Turner and Newman (2005) reported a distinct absence of inositol phosphates in wetland soils and benthic floccu-

A further consideration in the selection of an extraction scheme is compatibility with the detection method to be used. Cade-Menun and Preston (1996) reported that extraction with 0.25 M NaOH and 0.05 M EDTA is suitable for both the extraction of organic phosphorus and subsequent characterization by solution 31P nuclear magnetic resonance (NMR) spectroscopy. Similar extraction conditions have been used in the extraction and determination of inositol phosphates from soils (Turner and Richardson, 2004) and manures using 31P NMR (Turner and Leytem, 2004). Given the ubiquity of inositol phosphates in soils and their known physicochemical behaviour, it is reasonable to suppose that high concentrations would be found in sediments, and that once buried, they would remain bound as metal precipitates. However, a study of the distribution of inositol phosphates in coastal marine and estuarine sediments from Tokyo Bay showed that although riverine suspended particulate matter and estuarine sediments contained appreciable amounts of inositol phosphates, the concentrations decreased progressively towards the mouth of the bay (Suzumura and Kamatani, 1995b). Further, inositol phosphate concentrations were high in surface sediments, but were almost completely absent in deeper layers (Fig. 16.4).

Inorganic, organic P (µmol/g ) IP6−P (⫻10−2 µmol/g ) 15

20

0

10

10

15

Inorganic

5

Organic

5

15

20

20

25

25

0

5

10

15

20

Inorganic

10

Organic

5

IP6

0

IP6

Depth (cm)

0

Fig. 16.4. Vertical distribution of inositol hexakisphosphate (IP6), inorganic phosphate and organic phosphorus from two cores in Tokyo Bay, Japan. (Reproduced from Suzumura and Kamatani, 1995b with permission from Elsevier.)

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lent from the Florida Everglades, USA, with the majority of the organic phosphorus being present as phosphate diesters. A possible explanation for this observation is that either there is no terrestrial source of myo-inositol hexakisphosphate to this system or the rates of decomposition and removal are much greater than the rate of supply from terrestrial sources. On the basis of these few examples, it is evident that inositol hexakisphosphate is not as refractory or immobile as previously thought, and that remobilization and mineralization of this organic phosphorus species may comprise an important source of bioavailable inorganic phosphate in sediment pore waters and the overlying water column.

Phosphorus Remobilization at the Sediment–water Interface While sediments are a major sink for both inorganic and organic phosphorus, there is strong interest in the biological, physical and chemical conditions that favour the release of the internal load of phosphorus from sediments back into the overlying water. The importance of the internal cycling and release of bioavailable inorganic phosphorus from sediments as a con-

tributor to algal blooms and eutrophication has long been appreciated. However, the emphasis has mostly been on bioavailable inorganic phosphate, and the role of organic phosphorus species in this process has largely been ignored. Phosphates in sediments may be either sorbed to, or co-precipitated with, metal hydroxyoxides, clay minerals and calcium carbonate, or bound to humic substances. In oxygenated waters, the sediments are covered with an oxidized microzone of iron(III) (e.g. Fe(OOH)) that will sorb phosphorus from overlying waters and act as a surface barrier, preventing diffusion of phosphorus from the sediment pores into the overlying water (Wetzel, 1999). This section summarizes the proposed mechanisms for sediment phosphorus remobilization. This process has two general components: (i) the release of phosphorus species from the particulate phase into the pore water, which in the case of organic phosphorus may involve either desorption or mineralization; and (ii) the transport of this phosphate-enriched pore water into the overlying water. The processes involved in both components is shown schematically in Fig. 16.5 (Boström et al., 1982; Wetzel, 1999; Golterman, 2001) and this section attempts to reconcile these with the observed behaviour of inositol phosphates.

Water TRANSPORT MECHANISMS

Diffusion

Wind-induced turbulence

Bioturbation

Gas ebullition

SEDIMENT

Dissolved phosphorus PHYSICO CHEMICAL MOBILIZATION

Desorption

Dissolution

Ligand exchange

Enzymatic MICROBIAL hydrolysis MOBILIZATION

Particulate phosphorus

Fig. 16.5. Schematic diagram showing important processes involved in the release of phosphorus from sediments. (Modified from Boström et al., 1982 and Wetzel, 1999.)

Inositol Phosphates in Aquatic Systems

Anaerobic conditions Phosphate is released from sediments under anaerobic conditions, and the most frequently advanced explanation for this behaviour is the solubilization of bound phosphates when iron(III) is reduced to soluble iron(II) (Einsele, 1936; Mortimer, 1941, 1942). Despite the popularity of this direct reduction or oxygen control model (Fig. 16.6), Golterman (2001) has argued that there is little evidence to support it. Instead, he and others have proposed that sulphate reduction under strongly reducing conditions would lead to the formation of insoluble FeS from Fe(OOH), thus indirectly releasing adsorbed phosphate species from Fe(OOH)≈P (Fig. 16.6). Several studies provide strong evidence in support of this mechanism for phosphate release (Caraco et al., 1989; Roden and Edmonds, 1997; Rozan et al., 2002). Given that myo-inositol hexakisphosphate is adsorbed to Fe(OOH) in preference to phosphate (Celi et al., 2001), its release from anoxic sediments by this mechanism is perhaps more feasible than that of solubilization of Fe(OOH), especially as iron(II)-phytate is reportedly insoluble (De Groot and Golterman, 1993). Golterman et al. (1998) also suggested that anaerobic fermentation of phytate in sediments is a possible source

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of phosphate release, which might explain the dramatic decrease in myo-inositol hexakisphosphate in anaerobic sediments of Tokyo Bay (Suzumura and Kamatani, 1995b) and the absence of inositol phosphates in Florida wetlands (Turner and Newman, 2005).

Aerobic conditions Phosphate is released from sediments under aerobic conditions, especially in shallow, non-stratified systems that are well oxygenated (Boström et al., 1982). Bacterial mineralization of organic phosphorus through hydrolysis of phosphate esters by enzymes such as alkaline phosphatase is thought to be an important remobilization mechanism (Fig. 16.7). These extracellular phosphohydrolytic enzymes are produced by algae and bacteria and are reported to have high activity in the suspended particulate and sediment phases (Boon, 1989). However, inositol hexakisphosphate is not amenable to hydrolysis by alkaline phosphatase (McKelvie et al., 1995) and many algae, while possessing phosphomono- and diesterase activity, show no phytase activity (Whitton et al., 1990, 1991). Consequently, inositol phosphate

Fig. 16.6. Possible mechanisms for the release of phosphorus species from anaerobic sediments in response to sulphate reduction and direct reduction of iron(III). Diss. P = dissolved phosphorus; SRB = sulphate-reducing bacteria; IRB = iron-reducing bacteria. Fe(OOH)˜P represents iron-associated phosphorus species (phosphate, organic phosphorus). (Modified from Roden and Edmonds, 1997.)

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Fig. 16.7. Possible mechanisms for the release of phosphorus species from sediments in aerobic, shallow water systems. The scale of the arrows is not representative of flux magnitude.

species have been considered bio-unavailable. However, some cyanobacteria such as Calothrix exhibit phytase activity (Livingstone et al., 1982, 1983), which may represent an important mechanism for the hydrolysis of myo-inositol hexakisphosphate. Similarly, a number of other microorganisms in aquatic environments have the capacity to utilize phytate (see Hill and Richardson, Chapter 5, this volume). Organic phosphorus mineralization may also occur under aerobic sediments due to bacterial respiration (Fig. 16.7). Oxygen is used as an electron acceptor in this process, although nitrate may also be utilized as oxygen becomes depleted. Release of inorganic phosphate under net aerobic conditions has been reported, but it is suggested that even at moderately oxidizing redox potentials anaerobic microzones will occur on the sediment surface and that either direct or indirect reduction can occur at these sites (Boström et al., 1982). In shallow water bodies that are well lit, sediments may be covered by benthic algal films. Photosynthetic production of oxygen by this micro-phytobenthos will increase the thickness of the oxidized microzone and reduce the flux of phosphorus from pore waters. They may act as a physical barrier that retards upwards diffusion of pore water, although they can enhance the uptake of phosphorus from overlying waters into the sediments (Underwood, 2001). In the dark,

benthic algal films may also release phosphorus from sediment due to respiratory breakdown of organic phosphorus (Graneli and Sundback, 1985). Given the complexity of these microphytobenthos assemblages, it is not improbable that they might produce phytase as a means of utilizing myo-inositol hexakisphosphate and other organic phosphorus substrates, although this has yet to be tested.

pH and ligand exhange processes As sediments become anoxic their pH decreases due to the increase in dissolved carbon dioxide. In eutrophic hardwater systems this can solubilize apatite and release associated phosphate (Golterman, 1998). On the other hand, increasing the pH decreases the sorption capacity of iron(III) hydroxyoxides, and hence the amount of phosphate or organic phosphorus adsorbed. This behaviour may be due to ligand competition by hydroxyl groups and two possible mechanisms have been proposed (Fig. 16.8). Sediment release of inositol hexakisphosphate by this mechanism is quite feasible given that sodium hydroxide is successfully used to extract myo-inositol hexakisphosphate from soils and sediments. Data from Rippey (1977) cited in Boström et al. (1982) show that higher rates of phosphorus release from

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273

O

O Me — O— P — O — R + OH

−

+

Me — O — H

−O —

OH

P —O —R OH

(a)

Fe — O

O R

−O

Fe — OH

+ Fe — OH

−O

O P

—

+

P

OH−

—

O

—

Fe — O

O—R

(b) Fig. 16.8. Possible hydroxyl ligand exchange mechanisms that would account for phosphorus removal at higher pH (a) from Lijklema (1977) and (b) from Andersen (1975) reported in Boström et al. (1982). R represents either a proton or an organic moiety.

Physical and biological sediment perturbation

sediments occurred from about pH 8.2 upwards. This corresponds to the pH of sea water, and suggests that ligand exchange associated with increasing pH in Tokyo Bay might explain some of the loss of myo-inositol hexakisphosphate from marine sediments there. However, the increasingly negative charge on inositol hexakisphosphate (~ −9 at pH 8.2) will tend to oppose, if not entirely negate, ligand exchange as a possible release mechanism for myo-inositol hexakisphosphate. Bacteria from lake sediments are also reported to produce organic acids in conjunction with carbohydrate metabolism and growth in aerobic systems. These organic acids can sequester metal ions in metal–phosphate complexes (e.g. FePO4) resulting in the solubilization of the phosphorus species (Boström et al., 1982).

In shallow waters, wind- and tidal-induced turbulence will cause suspension of surficial sediment, which favours release of phosphorus-rich pore water back into the water column over the much slower diffusion process. However, phosphorus mobilized in this manner may be rapidly readsorbed to suspended particulate matter (Holdren and Armstrong, 1980). Other physical processes capable of disturbing the sediment include gas ebullition (e.g. nitrogen gas as part of denitrification). The sediment surface layer may also be disrupted through bioturbation and bioirrigation by organisms such as tubificid worms, chironomids and benthivorous fish.

Salinity

Future Research

Changes in salinity have been observed to release both inorganic and organic phosphorus from sediments, most probably through a combination of the lysis of bacterial cells and ligand exchange (Gardolinski et al., 2004). It was noted that significant organic and inorganic phosphorus release (10 µg P/l) occurred at salinity values >10 on the practical salinity scale, and that this was followed by rapid hydrolysis and release of bioavailable reactive phosphate. The potential for inositol phosphate remobilization by this route is still unknown.

There is little appreciation of the magnitude of inositol phosphate transport within the aquatic environment. Given that inositol phosphates can constitute a sizeable fraction of the total phosphorus in soil particles, riverine transport of suspended sediments is a potentially important source of phosphorus in estuaries and coastal waters. This is exemplified using data from the UK. Taking 133 mg P/kg as an average concentration of myo- and scyllo-inositol hexakisphosphates for soils (Turner et al., 2003, 2005b) and

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an annual suspended solids load for UK rivers of 45 × 109 kg (Littlewood and Marsh, 2005), an estimated annual load of 6 million kg of inositol hexakisphosphate-phosphorus can be calculated. This represents a relatively minor amount of phosphorus compared with the equivalent annual load of dissolved phosphate of ~370 million kg (Littlewood and Marsh, 2005). However, as the dissolved phosphate load includes a considerable contribution from point sources (e.g. Sharpley and Withers, 1994), such as sewage and industry, the inositol phosphate load probably accounts for a substantial component of the phosphorus export from diffuse sources. Transport of inositol phosphates to aquatic ecosystems is of potential significance, because it is becoming increasingly evident that they are not refractory, immobile phosphorus species as was once thought. The detection of myo-inositol hexakisphosphate in the pore and overlying waters suggests that there is diffusional transport from sediments, and that the released compounds may be converted to more bioavailable forms in the presence of hydrolytic enzymes. The literature abounds with reference to the association of inositol phosphate with higher-molecular weight organic matter such as humic material, but as yet there is no clear understanding of the nature of these interactions or their importance in the cycling of inositol phosphates. To a large extent, the study of inositol phosphates in aquatic systems has been hampered by the absence of suitable and accessible techniques for their analysis and detection (Turner et al., 2002b). For example, it is unclear whether myoinositol hexakisphosphate detected in overlying

waters is present in true solution or a colloidal form. This question will probably not be answered by measurements based on gel filtration or ion exchange after hypobromite oxidation, and the use of less invasive preparation and separation techniques will be required. Similarly, studying the role of phytase in the hydrolysis of inositol phosphates in sediments and waters has been complicated by the lack of an artificial substrate that would allow straightforward measurement of phytase activity (Turner et al., 2002b). The recent development of a `tethered’ inositol phosphate compound that can be used as a substrate in activity measurements assists greatly in elucidating the role of bacteria and algae in hydrolysing myo-inositol hexakisphosphate (Berry and Berry, 2005). Further elucidation of the behaviour of myoinositol hexakisphosphate at the sediment–water interface will also require improvement of sampling techniques other than that offered by the straightforward collection of sediment cores and pore waters. Devices such as benthic chambers for flux measurements across the sediment–water interface, and diffusive gradients in thin films (DGT) (Zhang et al., 1998) with binding phases specifically designed for organic phosphorus, would provide valuable information in this respect.

Acknowledgement I am indebted to Dr Philippe Monbet for his critical comments and assistance in redrafting some diagrams.

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Sommers, L.E., Harris, R.F., Williams, J.D.H., Armstrong, D.E. and Syers, J.K. (1972) Fractionation of organic phosphorus in lake sediments. Soil Science Society of America Proceedings 36, 51–54. Stephens, L.R. and Irvine, R.F. (1990) Stepwise phosphorylation of myo-inositol leading to myo-inositol hexakisphosphate in Dictyostelium. Nature 346, 580–583. Suzumura, M. and Kamatani, A. (1993) Isolation and determination of inositol hexaphosphate in sediments from Tokyo Bay. Geochimica et Cosmochimica Acta 57, 2197–2202. Suzumura, M. and Kamatani, A. (1995a) Mineralization of inositol hexaphosphate in aerobic and anaerobic marine sediments: implications for the phosphorus cycle. Geochimica et Cosmochimica Acta 59, 1021–1026. Suzumura, M. and Kamatani, A. (1995b) Origin and distribution of inositol hexaphosphate in estuarine and coastal sediments. Limnology and Oceanography 40, 1254–1261. Turner, B.L. (2005) Organic phosphorus transfer from terrestrial to aquatic environments. In: Turner, B.L., Frossard, E. and Baldwin, D.S. (eds) Organic Phosphorus in the Environment. CAB International, Wallingford, UK, pp. 269–294. Turner, B.L. and Leytem, A.B. (2004) Phosphorus compounds in sequential extracts of animal manures: chemical speciation and a novel fractionation procedure. Environmental Science and Technology 38, 6101–6108. Turner, B.L. and Newman, S. (2005) Phosphorus compounds in wetland soils: the importance of phosphate diesters. Journal of Environmental Quality 34, 1921–1927. Turner, B.L. and Richardson, A.E. (2004) Identification of scyllo-inositol phosphates in soil by solution phosphorus31 nuclear magnetic resonance spectroscopy. Soil Science Society of America Journal 68, 802–808. Turner, B.L., McKelvie, I.D. and Haygarth, P.M. (2002a) Characterisation of water-extractable soil organic phosphorus by phosphatase hydrolysis. Soil Biology and Biochemistry 34, 27–35. Turner, B.L., Papha´zy, M.J., Haygarth, P.M. and McKelvie, I.D. (2002b) Inositol phosphates in the environment. Philosophical Transactions of the Royal Society, London, Series B 357, 449–469. Turner, B.L., Mathieu, N. and Condron, L.M. (2003) Quantification of myo-inositol hexakisphosphate in alkaline soil extracts by solution 31P NMR spectroscopy and spectral deconvolution. Soil Science 168, 469–478. Turner, B.L., Cade-Menun, B. J., Condron, L.M. and Newman, S. (2005a) Extraction of soil organic phosphorus. Talanta 66, 294–306. Turner, B.L., Mathieu, N., Condron, L.M. and Chen, C.R. (2005b) Quantification and bioavailability of scylloinositol hexakisphosphate in pasture soils. Soil Biology and Biochemistry 37, 2155–2158. Underwood, G. J.C. (2001) Microphytobenthos. In: Steele, J.H. (ed.) Encyclopedia of Ocean Sciences. Elsevier Science, Oxford, pp. 1770–1777. Valaja, J., Plaami, S. and Siljander-Rasi, H. (1998) Effect of microbial phytase on digestibility and utilisation of phosphorus and protein in pigs fed wet barley protein with fibre. Animal Feed Science and Technology 72, 221–233. Weimer, W.C. and Armstrong, D.E. (1977) Determination of inositol phosphate esters in lake sediments. Analytica Chimica Acta 94, 35–47. Weimer, W.C. and Armstrong, D.E. (1979) Naturally occurring organic phosphorus compounds in aquatic plants. Environmental Science and Technology 13, 826–829. Wetzel, R.G. (1999) Organic phosphorus mineralization in soils and sediments. In: Reddy, K.R., O’Connor, G.R. and Schelske, C.L. (eds) Phosphorus Biogeochemistry in Subtropical Ecosystems. Lewis Publishers, Boca Raton, Florida, pp. 225–245. White, R.H. and Miller, S.L. (1976) Inositol isomers: occurrence in marine sediments. Science 193, 885–556. Whitton, B.A., Potts, M., Simon, J.W. and Grainger, S.L. J. (1990) Phosphatase activity of the blue-green alga (cyanobacterium) Nostoc commune UTEX 584. Phycologia 29, 139–145. Whitton, B.A., Grainger, S.L. J., Hawley, G.R.W. and Simon, J.W. (1991) Cell-bound and extracellular phosphatase activities of cyanobacterial isolates. Microbial Ecology 21, 85–98. Zhang, H., Davison, W., Gadi, R. and Kobayashi, T. (1998) In-situ measurement of dissolved phosphorus in natural waters using DGT. Analytica Chimica Acta 370, 29–38.

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Index

Abiotic reactions of inositol phosphates in aquatic systems 263–264 in soil 207–217 see also Adsorption Acid dissociation constants (pKa) 17–18, 214, 215 Acid hydrolysis, soil extracts 41, 42, 49 Acid phosphatases 244 see also Purple acid phosphatases Acid protease 137–138 Adenosine diphosphate–glucose pyrophosphorylase (AGP) 119 Adsorption (sorption) inositol phosphates in aquatic systems 263–264 inositol phosphates in soils 207–213 effects on surface properties 216–217 effects on utilization by plants 245 role of calcium carbonate, clays and organic matter 211–213 role of iron and aluminium oxides 208–211 solution characteristics affecting 214–216 phytases in soils 223–225 Aerobic conditions, sediments 271–272 Agranoff’s turtle 4, 5 Algal films, benthic 272 Alkaline phosphatase 271 Alkaline phytases 102, 103 Alum (aluminium sulphate) 163 Aluminium ions inositol phosphate hydrolysis and 175, 177–178 inositol phosphate reactivity 172–173 Aluminium oxides, soil adsorption and 208–211 Amoebae 195 Anaerobic conditions in aquatic systems 269, 271 inositol phosphate hydrolysis 192, 269–270

phytase synthesis 83, 84, 85 phytate-degrading microorganisms 65, 66 soil inositol phosphate content 192 Analytical separation methods 24–29 Animal feeds see Feeds, animal Animal manures see Manures, animal Animal nutrition 133–143 see also Dietary manipulation Anions 174, 225, 231–232 appA-encoded phosphatase see Escherichia coli, AppA phytase Aquaculture 122 Aquatic systems 261–274 amounts of inositol phosphates 264–270 phosphorus remobilization at sediment–water interface 270–273 physicochemistry of inositol phosphates 262–264 sources of inositol phosphates 261–262 see also Water bodies Arabidopsis purple acid phosphatases 244 seed phytic acid biosynthesis 114, 115–116 transgenic phytase-expressing 102, 104–105, 246 vacuolar adenosine triphosphatase 117 Arbuscular mycorrhizae 243, 248–250, 252 Aspergillus expression systems 141 Aspergillus fumigatus phytase 99, 101, 138, 140, 141, 222 Aspergillus niger (formerly A. ficuum) phytases (mainly PhyA) 64, 67, 80, 82 in animal feeds 135–136, 173 Apase6 104, 105 applications 68, 102 catalytic mechanisms 98, 99–100 disulphide bridges 101 expression systems 68, 100, 141 glycosylation 100, 101, 142 279

280

Index

Aspergillus niger (Continued ) improving pH profile/catalytic properties 140–141 PhyB 100 properties 67–68, 69, 223 proteolysis resistance 139 soil interactions 223, 225, 226–227, 228, 229, 230–231 transgenic plants overexpressing 138, 246, 247, 252–253 Aspergillus oryzae phytase 222 Aspergillus terreus phytase 69, 98 Axenic culture 245

Bacillus spp. phytases 64–65, 66, 67, 68, 89 applications 103 catalytic mechanism 102–103 degradation pathways 79, 81 expression systems 68, 141–142 inoculation of plants 245 properties 69, 222 regulation of synthesis 82, 83 Baculovirus expression system 142 Barley low phytate (lpa) varieties 113, 118–119, 126 animal feeding studies 122, 138, 155, 156 human feeding studies 124, 126 phytases 80, 87 seeds 14, 15, 111 Barrel medic (Medicago truncatula) phytase 104–105, 246 β-propellor phytases (BPP) 61, 71, 79, 102–103, 106 Biofarming 102 Bis-diphospho-myo-inositol tetrakisphosphate (IP8 ) 3, 114, 116 Burkholderia spp. 70, 245

Calcareous soils 159, 160–161, 193, 211 Calcite 211, 263 Calcium bioavailability, low-phytate crops 121, 122, 124, 125 faecal excretion 135 Calcium carbonate 212 Calcium ions effects on adsorption 216, 263 inositol phosphate reactivity 171–172 phytase activity and 82–83, 102, 103, 231 Calcium/phosphorus ratio, dietary 136, 137 Calcium phytate 171, 172 adsorption in soil 211 in culture media 62, 63 precipitation in soils 213 utilization by plants 247–248, 249 cAMP receptor protein (CRP) 82, 84 Capillary electrophoresis 27–29

Carbohydrases 137 Carbon content, animal manures 159, 160, 161, 162, 164 source, phytase synthesis and 82, 84–85 starvation, phytase synthesis and 81, 82, 83, 84 Carbon-14 (14C) labelling studies 50–53, 56–58, 199, 200 Cations adsorption in soils and 215–216 complexation in soils 213 inositol phosphate hydrolysis and 175 inositol phosphate reactivity 171–173 phytase activation/inhibition in soil 231 Cattail (Typha spp.) 79 Cattle 70, 126, 173 Caulobacter crescentus 88 Cellulase 137–138 Cereal grains 111–112 Chelating agents 173–174, 232 Chemical degradation 7, 8 Chilton Conference on Inositol and Phosphoinositides (1984) 1 Citrate 63, 134, 136–137, 232 Citrobacter braakii phytase 66, 67, 69, 139 Clays adsorption of phytases 223, 224, 228–229 retention of organic phosphorus 207, 208 sorption of inositol phosphates 212, 263 Clover, subterranean (Trifolium subterraneum) 246, 247, 251, 252 Conformational inversion 8–10, 11 Conformational isomers 8, 9 Conformers 2 Consensus-1 phytase 139, 141 Consensus-7 phytase 141 Copper chelate stability 173–174 Cultivated soils 192, 193 Culture media 62–63, 66–67 Cyclic adenosine monophosphate (cAMP) 82, 84 1,2-Cyclohexanediamine tetraacetate (CDTA) 174, 175, 176, 177–178 Cysteine phosphatases 61, 103–104 Cysteine phytases (CPhy) 104, 106

Denaturing gradient gel electrophoresis (DGGE) 71 Desorption, inositol phosphates 213–214, 271 Diet, phosphorus composition of manures and 153, 154 Dietary manipulation environmental fate of manure phosphorus and 161–163 future research needs 164–165 overall benefits 164

Index

phosphorus content of manure 126, 133–134, 135, 153–161 see also Feeds, animal Diethylene triaminepentaacetate (DTPA) 175, 176, 177–178 Diphospho-myo-inositol pentakisphosphate (IP 7) 3, 114, 116 Diphospho-myo-inositol tetrakisphosphate (PP-IP4) 3 Drying, soil, phytase activity and 233 Duckweed 262

Ecology, phytate-degrading microorganisms 68–71 Ectomycorrhizal fungi 243, 248–250, 251–252 EDTA see Ethylenediaminetetraacetate Electrospray ionization (ESI) 23, 29–30 Electrospray ionization–time of flight–mass spectrometry (ESI–TOF–MS) 23–24, 29–38 direct 31–34 principles 29–30 size-exclusion chromatography with 34–37, 38 Enterobacter 66, 89 Enterobacter cloacae 78, 79, 80 Enterobacteriaceae 90 Environmental fate, manure phosphorus 161–163, 262–263 Environmental issues 111–112, 133, 150–165 Environmental samples high-performance chromatography and mass spectrometry 25, 34, 38 NMR spectroscopy 18–19 Epimerization 2 microbial, in soils 49–50, 55–58 reactions 199 Epimers 2 Erosion, soil 161 Escherichia coli agp-encoded acid phosphatase 66, 83, 88 AppA phytase 66, 67, 83–85, 88, 134–136 expression systems 141 with improved thermostability 139 improving pH profile/catalytic properties 140 proteolysis resistance 139–140 expression systems 68, 102, 141 phytases 64, 65–67, 78, 91 catalytic mechanism 98, 99 degradation pathways 79, 80 disulphide bridges 101 expression systems 68, 100, 138, 141 glycosylation 138, 139 in vivo function 88–89, 90 properties 69, 222 regulation of synthesis 81, 82, 83–85 Ethylenediaminetetraacetate (EDTA) -exchangeable phytase-hydrolysable phosphorus 179–180

281

extraction methods 152–153, 190, 191 ligand exchange studies 174, 175, 176, 177–178 Exchange spectroscopy (EXSY), random delay 10, 19 Extraction methods animal manures 152–153, 170–171 aquatic systems 263, 266–269 soil 41–42, 190

Feeds, animal pelleting 138 phytase supplements see under Phytases phytate content 111–112, 133 using low-phytate crops see Low-phytate crops see also Dietary manipulation Fermentation technologies, phytase production 67 Ferrihydrite 208, 209, 210, 213 Fertilizers, phosphorus-based 102, 242 effects on soil inositol phosphates 193 phytase-expressing transgenic plants 252 Fish, diets using low-phytate crops 122 Flow-scintillation analysis, isotopic tracers in soils 54–55, 56 Food, applications of phytases 102 Forest soils 192 Formic acid 84, 136–137 Fractionation, phosphorus in animal manures 152, 170 Fragmentation, mass spectral ion inositol phosphates 32–34, 35, 37, 38 inositol stereoisomers and products 43–46 Freezing, phytase denaturation 233 Fungal phytases 65, 67, 82, 99–100, 222 applications 102 consensus constructs 139, 141 disulphide bridges 101 glycosylation 100, 101, 142 transgenic plants overexpressing 246–247

Gas chromatography 24 Gas–liquid chromatography 24 Gel chromatography 25 Germination, phytate degradation 87 Gibberellic acid 87 Globoids 111, 116–117 Glossary of terms 2–5 Glucose, effects on phytase synthesis 82 Glucose-1-phosphatase 78, 83 Glycosylation Escherichia coli phytase 138, 139 fungal phytases 100, 101, 142 phytases in soil 230 Goethite (Fe(OOH)) adsorption to 209, 210, 211, 214–215, 263 desorption from 213, 271 Grassland soils 192, 195

282

Index

Hebeloma cylindrosporum phytase 225, 228 Hedley fractionation method 152, 170 Heteronuclear multiple quantum correlation plus twodimensional total correlation spectroscopy (HMQC-TOCSY) 15–17, 20 High-performance liquid chromatography (HPLC) 24, 54 reversed-phase (RP-HPLC) 23, 25, 26, 27 High-performance size-exclusion chromatography (HP-SEC) 23, 34–37 mass fragmentometry approach 37, 38 principle and technique 34–36 selected ion monitoring (SIM) approach 36–37, 38 Histidine acid phosphatases (HAP) 61, 98–102 disulphide bridges 101 glycosylation 100 phytate degradation pathway 79 substrate specificity site 99–100 Histidine acid phytases (HAPhy) 99–102, 106 Human nutrition 112 low-phytate crops 122–126, 127–128 phytase supplements 135 Humic materials 191, 212–213, 223–224, 228 Hydrogen-bonding interactions 18 Hypobromite oxidation (alkaline bromination) 190, 191, 196–197, 264, 265, 267, 268

Identification of inositol phosphates in aquatic systems 266–269 by mass spectrometry see Mass spectrometry by NMR spectroscopy 7–20 in environmental samples 18–19 in impure samples 15–17 in a mixture without separation 12–14 in plant seeds 14 purified compounds 10–12 in soils 18–19, 46, 48, 49, 186, 190, 191 Inductively coupled plasma (ICP) mass spectrometry (ICP–MS) 23, 27 Infrared spectroscopy 46, 47 Inositol 2 allo-Inositol 3, 198, 199 chiro-Inositol infrared spectroscopy 46, 47 proton NMR spectroscopy 46, 48 in soil 57 D-chiro-(+)-Inositol 3, 194 mass spectrometry 43, 44 origins 49–50, 55, 57, 58, 198, 199 in soils 49–50, 194 identification 46–49 isotopic studies 50, 53, 54 L-chiro-(−)-Inositol 3, 198, 199 cis-Inositol 3, 198, 199 epi-Inositol 3, 198, 199

muco-Inositol 3 identification in soils 47 origins 198, 199 myo-Inositol 2, 3, 194 Agranoff’s turtle 4 infrared spectroscopy 46, 47 mass spectrometry 43–46 NMR spectroscopy 16–17, 46, 48 origins 198, 199 in soil 50, 194 carbon and phosphorus pathways 55 identification methods 43–46, 47, 48 isotopic studies 50–53, 54, 56–58 microbial epimerization 49–50, 55–58 neo-Inositol 3, 194 origins 57, 198, 199 in soils 49, 50 scyllo-Inositol 3, 5, 194 origins 57, 198, 199 in soils 49, 50, 194 myo-Inositol bisphosphate 3, 32, 36, 37 D-chiro-(+)-Inositol hexakisphosphate 195, 196, 197 muco-Inositol hexakisphosphate 49, 195, 198, 199 myo-Inositol hexakisphosphate (InsP6) 2, 3, 4, 97 see also phytic acid in animal manures 150, 151–152, 153, 154, 170 dietary manipulation 153–158 environmental fate 162, 163 ligand effects 174–180 phosphorus solubility in soil and 159–161 reactivity with polyvalent cations 173 in aquatic systems 261–274 biosynthesis in seeds 114–116 cation complexes 171–173 ligand exchange 174–178 pH effects 172–173 in ruminant excreta 173 in soils 213, 214 stability calculations 174 conformational inversion 8–9, 10, 11 identification by NMR spectroscopy 14, 16–17, 190, 191 identification of hydrolysis products 13–14 mass spectrometry 32–33, 34 phosphorylases see Phytases pKa values 18, 214, 215 salts see Phytate size-exclusion chromatography and mass spectrometry 36–37, 38 in soils 187–188, 189, 193, 194 adsorption 209, 210–211, 212, 214–217 complexation 213, 214 desorption 213, 214 isotopic studies 50–53 origins 199

Index

neo-Inositol hexakisphosphate 195, 196, 197 scyllo-Inositol hexakisphosphate NMR spectroscopy 18–19, 190, 191 origins 195, 199 potential function 201 in soils 194–197 Inositol hexakisphosphates origins 195–199 in soils 187, 188–189, 196–197 myo-Inositol kinase 114, 115 myo-Inositol monophosphate 3 mass spectrometry 32, 33, 34 NMR spectroscopy 12, 16 size-exclusion chromatography and mass spectrometry 36–37 D-myo-Inositol 3-monophosphate synthase (MIPS) 114–115, 120 myo-Inositol pentakisphosphate 3 biosynthesis in seeds 114, 116 conformational inversion 10, 11 mass spectrometry 32 NMR spectroscopy 14 Inositol pentakisphosphates, in soil 187–190 Inositol phosphates 2, 3 in animal manures 150–165, 195–199 in animal nutrition 133–143 in aquatic systems 261–274 identification see Identification of inositol phosphates nomenclature 1–5 plant utilization 242–254 separation and detection by mass spectrometry 23–38 in soil see under Soils D-chiro-(+)-Inositol phosphates 195, 198 myo-Inositol phosphates 3 in aquatic systems 261–262 in low phytic acid seeds 112–113 metabolic pathways in seeds 113–116 origins 198 in soils 194 neo-Inositol phosphates 194, 195, 198 scyllo-Inositol phosphates origins 195, 198 in soils 194–195 myo-Inositol polyphosphate 2-kinase 114, 116 Inositol stereoisomers (and phosphorylated derivatives) 3, 5 in aquatic systems 262, 269 in soils 41–58, 193–201 myo-Inositol tetrakisphosphate 3 conformational inversion 10, 11 mass spectrometry 32 NMR spectroscopy 14 myo-Inositol trisphosphate 3, 8, 18, 32 myo-Inositol 1,3,4-trisphosphate 5/6-kinase 114, 115, 116

283

myo-Inositol 1,4,5-trisphosphate 3/6-kinase 114, 115, 116 Inosose, mass spectrometry 43–46 DL-epi-Inosose, mass spectrometry 43, 44 International Union of Pure and Applied Chemistry (IUPAC) and International Union of Biochemistry (IUB) 1, 4–5, 7, 78 Ion chromatography 26–27, 28 Ion-exchange chromatography 24, 26 radionuclide tracers in soils 54, 56 Ion-pairing reversed-phase high-performance liquid chromatography (HP-ion pair-RPLC) see under High performance liquid chromatography 23, 25, 27 Iron bioavailability dietary phytase supplements 135 low-phytate crops 123–124 Iron hydroxide, addition to soil 180, 181 Iron oxides adsorption to 208–211, 263 desorption from 213–214, 271 Iron(II) phytate, in aquatic systems 271 Iron(III) ions chelate stability 173–174 inositol phosphate hydrolysis and 177–178 inositol phosphate reactivity 171–172, 172 sorption in aquatic systems and 263, 270, 271 Iron(III) phytate, in soils 50–53, 192, 213 Isoelectric points, phytases 222, 225, 227 Isomers conformational 8, 9 positional 4–5 Isotopic tracer studies, in soils 50–58, 199, 200, 248, 252–253

Kaolinite 208, 209–210, 212, 228–229 Klebsiella spp. phytases 66, 69, 89 purification and characterization 64, 65 regulation of synthesis 82

Lactic acid 136–137, 138 Lactic acid bacteria (Lactobacillus spp.) 66, 69, 88, 138, 141–142 Legumes 111, 112 Ligands based fractionation assay 178–180 exchange processes 174–178, 272–273 sources 174 stability of cation complexes 173–174 Light radiation, phytase degradation 234 Lily (Lilium longiflorum), phytate-degrading enzymes 79, 81, 87, 103 Low-phytate crops 112–113, 117–128 animal feeding 120–122, 155, 156

284

Index

Low-phytate crops (Continued ) combined with phytase supplements 158, 159, 163 environmental fate of manure phosphorus 161–162 faecal phosphorus excretion 121, 122, 135 human nutrition 122–126 seed phosphorus and ruminant nutrition and 126–127 low phytic acid (lpa) genotypes 112–113, 114, 115–116, 117–119 Lucerne 102, 252 Lupin (Lupinus spp.) phytases 79, 80, 244, 246 phytate-degrading microorganisms 70 utilization of soil phytates 247–248

Magnesium, in low-phytate rice 125–126 Maize low phytate (lpa) varieties 112, 113, 117, 118, 119 animal feeding studies 120–121, 122, 155 human nutrition studies 123–125 phytase supplements with 138 seed phytic acid biosynthesis 114, 115–116 phytases 87, 98, 246 seeds 111, 112 utilization of soil phytate 248 Manures, animal 150–165 carbon/phosphorus ratios 159, 160, 161, 162, 164 environmental issues 111–112, 133, 150–165 inositol phosphates 150–165, 195–199 inositol phosphates reaching aquatic systems 262–263 nitrogen/phosphorus ratios 151, 153 phosphorus 150–165 analytical methods 151–153, 170–171 composition 151–153, 154 dietary manipulation 126, 133–134, 135, 153–161 environmental fate and dietary manipulations 161–163 phosphorus solubility in soil and 158–161 storage effects 158, 164 temporal changes in biological availability 180, 181 solubility and release of inositol phosphates 169–181 analytical methods 169–171, 178–180 characterizing relative stability 173–180 ligand exchange studies 174–178 reactivity with polyvalent cations 171–173 Mass fragmentometry 37, 38 Mass spectrometry (MS) 23–24, 29–38 capillary electrophoresis with 27 direct 31–34

electron-impact 43–46, 49 ion-pairing reversed-phase HPLC with 25, 27 radionuclide tracers in soils 54 size-exclusion chromatography with 34–37 Metal ions complexation of inositol phosphates 70, 71, 173–174, 213, 263 phytase inhibition 231 Microorganisms degradation of soil phytases 230–231 phosphorylated inositol stereoisomers 199, 201 phytases see Phytases, microbial phytate-utilizing see Phytate-degrading microorganisms soil, phosphorus utilization by plants and 243, 248, 251–252 synthesis of inositol phosphates 195, 199–201 Mineral nutrition, human 112, 123–126 Monogastric animals 70, 111–112 dietary manipulation of manure phosphorus 153–161 dietary phosphate supplements 153 dietary phytase supplements 156–158 diets using low-phytate crops 120–122, 155, 156 environmental fate of manure phosphorus 161–163, 262–263 phosphorus composition of manures 153, 154, 170 Montmorillonite 212, 224, 228–229 Mung bean phytase 79, 80, 87 Mycorrhizal fungi 243, 244, 248–250, 251–252

Nitrate, repression of phytase synthesis 85 Nomenclature 1–5 Non-ruminant animals see Monogastric animals Nuclear magnetic resonance (NMR) spectroscopy 7–20 acid dissociation constants 17–18 animal manures 152–153, 154 applications 9 aquatic system samples 269 conformational analysis 8–10 environmental samples 18–19 experimental details 19–20 intramolecular hydrogen bonding 18 plant root studies 251 protonation sequences at microscopic level 17–18 radionuclide tracers in soils 54–55 soil samples 18–19, 46, 48, 49, 186, 190, 191 solid-state 19 structural determinations 10–17 TOCSY technique 12–14, 15 Nutrient status regulation of phytase synthesis 88–89 soil inositol phosphates and 193, 195

Index

Organic matter, soil 212–213, 223–224, 228 Oxalic acid 175, 176, 177, 178, 232

Pantoea agglomerans 79, 80 Paper chromatography 24–25, 41–42 Paramecium 66, 79, 80, 201 Pathogenic infections 89–90, 103 Pelleting, animal feed 138 Penicillium simplicissimum 69 Peniophora lycii phytase 68, 140 interactions in soil 225, 227, 229, 230–231 properties 69, 222 pH inositol phosphate complexation and 213, 214 inositol phosphate hydrolysis and 172–173 optima of phytases 140–141, 229, 232 phosphorus mobilization in aquatic systems 272–273 phytase synthesis and 82 soil abiotic processes and 214–215, 225, 226 inositol phosphate content and 192–193 phytase activity and 232 PHO regulon 67, 83, 85–86 Phosphatases hydrolysis of soil organic phosphorus 243–244 see also Phytases Phosphate, inorganic animal feed supplements 111, 153 in animal manures 152, 173–180 fertilizers see Fertilizers, phosphorus-based inhibition of phytases in soil 231–232 ligand-exchangeable 179 microbial assimilation 88 regulation of phytate-degrading activity 87 seed 111, 112, 113, 117, 119 uptake by plants 243 see also Phosphorus Phosphatidylinositol (PtdIns) phosphates, in seeds 114, 115, 116, 117 Phosphoinositides 1 Phospholipase C 89–90 Phosphomonoesterases 78 see also phosphatases Phosphorus (P) in animal manures see under Manures, animal bioavailability animal feedstuffs 133–134 low-phytate crops 121, 122 dietary manipulation strategies see Dietary manipulation environmental issues 111–112, 133, 150–165 faecal excretion dietary phytase supplements and 135 low-phytate crops 121, 122, 135 manipulation strategies 133–134, 153–158

285

fertilizers see Fertilizers, phosphorus-based limitation/deficiency effects on plants 242, 243–245 inositol phosphate levels and 70, 193, 195 phytase synthesis and 66, 81, 83, 84, 85–86, 89 phytase-hydrolysable (PHP) see Phytasehydrolysable phosphorus seed 111, 112 crops with reduced total 126–127 in low-phytate crops 113, 119–120 non-ruminant nutrition studies 120–122 soil accumulation 151 run-off to water bodies 163, 164 solubility, after manure application 158–161, 164 utilization by plants 242–254 see also Inositol phosphates; Phosphate, inorganic Phosphorus-32 (32P) tracer studies 55, 58, 248 Phosphorus-33 (33P) tracer studies 56, 58, 252–253 Phthalic acid 175–176, 177, 178 Phytase-hydrolysable phosphorus (PHP) EDTA-exchangeable 179–180 ligand-based assay 178–180 in soils 193, 251, 252 Phytases (phytate-degrading enzymes) 4, 61, 78–91 animal feed supplementation 68, 102, 103, 133–143 activity in stored manures 136, 158 augmentation strategies 137–138 combined with low-phytate diets 158, 159, 163 constraints 138 determinants of efficacy 136–137 enhancing proteolysis resistance 139–140 environmental fate of manure phosphorus 162–163 impact on manure phosphorus 135, 156–158 improving pH profile/catalytic properties 140–141 nutritional impacts 134–135 production systems 141–142 site of activity in animals 135–136 thermostability 138–139 applications 102, 103 attributes and catalytic mechanisms 97–105 classification 61, 78–79, 97–98, 105, 106 in vivo function 87–90 ligand exchange effects 174–180 microbial 78 characterization of activity 63–65 degradation pathways 79, 80, 81 expression and production 65–67, 68 in vivo function 88–90

286

Phytases (Continued ) intra- and extracellular 65, 66, 89–90 ions and metabolites activating/inhibiting 231–232 properties 67–68, 69, 222 purification 63, 64 regulation of synthesis 79, 81–83 sources 65, 66 nomenclature 105, 106 phosphorylated stereoisomer hydrolysis 201 plant 78, 79 constitutive 86–87 degradation pathways 79, 80 germination-inducible 86, 87 in vivo function 88 ions and metabolites activating/inhibiting 231–232 regulation of synthesis 86–87 role in uptake of soil organic phosphorus 243–244, 245 plant roots 88, 244, 245–247 production systems 65–66, 67, 138, 141–142 regulation of synthesis 79–87 in soil 221–235 denaturation 232–233 factors affecting activity 222–223 ions and metabolites inhibiting/activating 231–232 microbial and proteolytic degradation 230–231 solid phase interactions 223–230 in soil–plant root environment 243–244 transgenic animals 221, 263 transgenic plants see Transgenic plants, phytaseexpressing 3-Phytases 78, 79 6-Phytases 78, 79 Phytate 2, 4 in animal feeds 111–112, 133 in animal manures 153 in human diet 112 phosphorus content 97 seed 111–128 utilization by plants 245, 247–248, 249, 250 see also myo-Inositol hexakisphosphate Phytate-degrading enzymes 4, 78 see also Phytases Phytate-degrading microorganisms 61–72, 250–251 in aquatic systems 271–272 assessment 61–65 ecology 68–71 inoculation of plants with 245 isolation case study 63 screening for 62–63 sources 65, 66 see also Phytases

Index

Phytic acid 4, 111 see also myo-inositol hexakisphosphate and phytate biosynthesis in seeds 113–116 Phytins 2, 4, 111 Phytoplankton 262 Pichia pastoris expression systems 68, 140, 141 Pigs see Swine Pinitol 198, 199 Pinus spp. 244, 250, 251 Pisolithus tinctorius 225 pKa values 17–18, 214, 215 Plants aquatic 262 in axenic culture 245 mycorrhizal associations 243 phytate-degrading enzymes see under Phytases synthesis of inositol stereoisomers 199–201 transgenic see Transgenic plants utilization of inositol phosphates 242–254 see also Rhizosphere; Roots; Seeds Pollen, phytate-degrading enzymes 79, 86–87 Polymerase chain reaction (PCR) 71 Positional isomers 4–5 Potassium, in low-phytate rice 125–126 Poultry 70, 97 dietary manipulation of manure phosphorus 155, 156–158, 159 dietary phytase supplements 134–135, 136–137, 156–158, 162–163 environmental fate of manure phosphorus 162–163, 262–263 low-phytate crop-based diets 120–121, 155, 156 low-phytate grains plus phytase supplements 158, 159 phosphorus composition of manures 153, 154 Precipitation, cation complexes in soils 214, 246 Preparative separations 24, 41–42 Proteases 137–138 Protein storage vacuoles (PSVs) 111, 116–117 Protein tyrosine phosphatase (PTP) 104 Proteolysis, phytases 139–140, 230–231 Protonation sequence, at microscopic level 17–18 Protozoa 201 see also Tetrahymena vorax and Parmecium Pseudomonas spp. phosphate utilization 89 phytases 69, 80, 82, 245 phytate degradation 62, 63, 66 Pteris vittata 202 Purple acid phosphatases (PAPs) 61, 104–105, 244, 246 Purple acid phytases (PAPhy) 104–105, 106 Pyrophosphatases 103

Quebrachitol, mass spectrometry 43, 44, 45, 46

Index

Radioisotope labelling studies 50–54, 199, 200, 248, 252–253 Raoultella terrigena phytase 79, 81, 82 Rhizopus oligosporus 66 Rhizosphere microorganisms 70, 88 phosphate uptake by plants 243, 248, 251, 252 phytases 224–225, 232, 244 Rice bran 82 low phytic acid (lpa) 113, 125–126 pathogens 90, 103 phytases 80, 104 seeds 111 Rivulariaceae 88 RNA polymerase, σs subunit 83–84 Roots phosphorus uptake from soils 243, 244–245 phytate-degrading enzymes 88, 244, 245–247 phytate-degrading microorganisms 70 see also Rhizosphere Rumen bacteria 89, 103–104 Ruminants 70, 112 low seed total phosphorus crops 126–127 myo-inositol hexakisphosphate in excreta 173 phosphorus composition of manures 153, 154, 170 Rye phytase 80, 87 Ryegrass Lolium perenne L. 248, 251–252

Saccharomyces carlsbergensis 201 Saccharomyces cerevisiae expression systems 68, 139–140, 141 phytases 69, 80, 91, 222 in vivo function 88 production and expression 65–66, 67 regulation of synthesis 85–86 Salinity, changes in 273 Salmonella dublin 90 Schwanniomyces castellii 69, 82 Second messengers 89–90 Sediments amounts of inositol phosphates 266–270 phosphorus remobilization mechanisms 270–273 physical and biological perturbation 273 sources of inositol phosphates 261–262 Seeds field emergence 120 inositol phosphates 112–113 NMR spectroscopy 14, 15 phosphorus see Phosphorus (P), seed phytate 111–128 deposition in globoids 116–117 lpa genotypes 112–113

287

metabolic pathways, genes and mutants 113–116 see also Low-phytate crops phytate-degrading enzymes 79, 86–87, 88 Selected ion monitoring (SIM) 33, 36–37, 38 Selenomonas ruminatum phytase 66, 82, 89, 103–104 Separation methods analytical 23, 24–29 preparative 24, 41–42 radionuclide tracers in soils 54 Sewage sludge 195 Shewanella oneidensis 71 Size-exclusion chromatography see High-performance size-exclusion chromatography Sodium hydroxide (NaOH) extractions 152–153, 170, 172, 190, 191 Sodium ions, inositol phosphate reactivity 171, Sodium phytate 171 in culture media 62, 63 utilization by plants 245, 247–248 Soils inositol phosphates 186–202 abiotic reactions 207–217 amounts 186–190 extraction and preparative chromatography 41–42 factors controlling amounts 190–193 NMR spectroscopy 18–19, 46, 48, 49, 186, 190, 191 reaching aquatic systems 262 inositol stereoisomers (and phosphorylated derivatives) 41–58, 193–201 isotopic studies 50–58 methodologies for characterizing 41–49 origins 49–50, 195–201 potential function 201 significance 49–50 iron hydroxide addition to manured 180, 181 ligand-based phytase-hydrolysable phosphorus assay 178–180 phosphorus accumulation 151 phosphorus solubility in manure-treated 158–161, 163, 164 phosphorus uptake by plants 243–245 phytases in see under Phytases phytate-degrading microorganisms 63, 65, 66, 70, 71, 250–251 phytate utilization by plants 245, 247–248, 249 Sorption see Adsorption Soybean low phytate varieties 113, 119, 120, 121 phytase (GmPhy) 79, 104, 105, 222, 246 seeds 111 transgenic phytase overexpressing 138 Spirodela polyrhiza 90 Stationary phase response 81, 83, 85, 89

288

Stereoisomers 5 see also Inositol stereoisomers Stress tolerance, low-phytate crops 118 Suillis phosphatases 225 Surface properties, effects of sorption 216–217 Swine dietary manipulation of manure phosphorus 155, 156, 158, 159 dietary phytase supplements 134–138, 156, 162–163 diets using low-phytate crops 122, 135, 155, 156, 162–163 environmental fate of manure phosphorus 162–163, 262–263 Escherichia coli phytase expression 99 low-phytate grains plus phytase supplements 158, 159 phosphorus composition of manures 153, 154

Temperature effects phytase stability 138, 232–233 phytase synthesis 82 Terminal-restriction fragment length polymorphisms (T-RFLP) 71 Terminology 1–5 Tetrahymena vorax 198, 201 Thermostable phytases 138–139 Time of flight (TOF) mass spectrometry 23–24, 30, 31–34 Tobacco (Nicotiana tabacum) 103, 246, 248, 249, 252 TOCSY see Two-dimensional total correlation spectroscopy Transgenic animals, phytase-expressing 221, 263 Transgenic plants, phytase-expressing 102, 103, 104–105, 221, 246–247 in animal feeds 138, 141 growth and phosphorus nutrition 248, 252–253 Trifolium subterraneum see Clover, subterranean

Index

Tritium (3H) tracer studies, in soil 56–58 Turtle (structure) 4, 5 Two-dimensional total correlation spectroscopy (TOCSY) 12–14, 15 heteronuclear multiple quantum correlation (HMQC) 15–17, 20 technique 19–20 Type III secretion systems 89–90

Ultraviolet (UV) absorbance detection 54

Vacuolar adenosine triphosphatase (V-ATPase) 116, 117 Vitamin D derivatives 134, 136

Water bodies amounts of inositol phosphates 264, 265, 266 transport of manure phosphorus to 151, 158–161, 163, 164, 262–263 see also Aquatic systems Water content of soil, phytase activity and 233 Wetland soils 192, 269–270 Wheat bran 67, 82 low phytic acid (lpa) genotypes 113, 126 phytase 79, 87, 245–246 seeds 111 utilization of phytates 248 Wolffiella floridana 262

Xanthomonas oryzae 90, 103

Yields, low-phytate crops 118–119

Zinc 124–125, 173–174