Inositol Phosphates: Linking Agriculture and the Environment

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Inositol Phosphates

LINKING AGRICULTURE

AND THE

ENVIRONMENT

This page intentionally left blank

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

CABI North American Office 875 Massachusetts Avenue 7th Floor Cambridge, MA 02139 USA Tel: +1 617 395 4056 Fax: +1 617 354 6875 E-mail: [email protected]

©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.

xi


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

4


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.


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;


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.


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.


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.


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


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


23

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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 (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).

214

L. Celi and E. Barberis

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


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.)

216


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


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.


219

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


221

222

T.S. George et al.

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

224

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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


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

225

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).

226

T.S. George et al.

(i) 100

(ii) 100

L.S.D. (P < 0.05)

L.S.D.(P < 0.05)

80

60 40 20 0 4 100

5

6

7

8

9

L.S.D. (P < 0.05)

80 60 40

Soil solid phase phytase activity (% added)

Soil solution phase phytase activity (% added)

80

40 20 0 4

5

6

7

8

9

100 L.S.D.(P < 0.05)

80 (b)

60 40 20

20 0

(a)

60

0 4

5

6

7

8

9

4

5

6

7

8

9

pH Solution

Solid 1h 48 h

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


(a) 12

Solution-phase phytase activity (nKat/g soil)

10 8 6 4 2 0 0 (b) 12

500

1000

1500

500 1000 Time (min)

1500

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).

228

T.S. George et al.

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


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

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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


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;

Relative phytase activity (%)

100

80

60

40

20

0 0

15

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Time (min)

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


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


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.

References Allison, S.D. and Vitousek, P.M. (2005) Responses of extracellular enzymes to simple and complex nutrient inputs. Soil Biology and Biochemistry 37, 937–944. Anderson, G. (1980) Assessing organic phosphorus in soil. 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. Baron, M.H., Revault, M., Servagent-Noinville, S., Abadie, J. and Quiquampoix, H. (1999) Chymotrypsin adsorption on montmorillonite: enzymatic activity and kinetic FTIR structural analysis. Journal of Colloid and Interface Science 214, 319–332. Basha, S.M. (1984) Purification and characterisation of an acid phosphatase from peanut (Arachis hypogaea) seed. Canadian Journal of Botany 62, 385–391. Bayan, M.R. and Eivazi, F. (1999) Selected enzyme activities as affected by free iron oxides and clay particle size. Communications in Soil Science and Plant Analysis 30, 1561–1571. Bianchetti, R. and Sartirana, M.L. (1967) The mechanisms of the repression by inorganic phosphate of phytase synthesis in the germinating wheat embryo. Biochimica et Biophysica Acta 145, 485–499. Brahms, W.H. and McLaren, A.D. (1974) Phosphatase reactions in columns of soil. Soil Biology and Biochemistry 6, 183–189. Brinch-Pedersen, H., Sørensen, L.D. and Holm, P.B. (2002) Engineering crop plants: getting a handle on phosphorus. Trends in Plant Science 7, 118–125. Brown, H.D. and Hasselberger, F.X. (1971) Matrix supported enzymes. In: Brown, H.D. (ed.) Chemistry of the Cell Interface. Academic Press, New York, pp. 185–258. Brugger, R., Nunes, C.S., Hug, D., Vogel, K., Guggenbuhl, P., Mascarello, F., Augem, S., Wyss, M., van Loon, A.P.G.M. and Pasamontes, L. (2004) Characteristics of fungal phytases from Aspergillus fumigatus and Sartorya fumigate. Applied Microbiology and Biotechnology 63, 383–389. Burns, R.G. (1982) Enzyme activity in soil: location and a possible role in microbial ecology. Soil Biology and Biochemistry 14, 423–427. Burns, R.G., Pukite, A.H. and McLaren, A.D. (1972) Concerning the location and persistence of soil urease. Soil Science Society of America Proceedings 36, 308–311. Casey, A. and Walsh, G. (2003) Purification and characterisation of extracellular phytase from Aspergillus niger ATCC 9142. Bioresource Technology 86, 183–188. Casucci, C., Okeke, B.C. and Frakenberger, W.T. Jr (2003) Effects of mercury on microbial biomass and enzyme activities in soil. Biological Trace Elements Research 94, 179–191. Chacon, N., Dezzeo, N. and Flores, S. (2005) Effects of particle-size distribution, soil organic carbon content and organic–mineral aluminium complexes on acid-phosphatases of seasonally flooded forest soils. Biology and Fertility of Soils 41, 69–72. Chadha, B.S., Harmeet, G., Mandeep, M., Saini, H.S. and Singh, N. (2004) Phytase production by the thermophilic fungus Rhizomucor pusillus. World Journal of Microbiology and Biotechnology 20, 105–109. Chang, C.W. (1966) Study of phytase and fluoride effects in germinating corn seeds. Cereal Chemistry 43, 129–142. Chang, R. and Schwimmer, S. (1977) Characterisation of phytase of beans (Phaseolus vulgaris). Journal of Food Biochemistry 1, 45–56. 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. Ciurli, S., Marzadori, C., Benini, S., Deiana, S. and Gessa, C. (1996) Urease from the soil bacterium Bacillus pasteurii: immobilisation on Ca-polygalacturonate. Soil Biology and Biochemistry 28, 811–817. Crook, E.M., Brocklehurst, K. and Wharton, C.W. (1970) Cellulose-insolubilised enzymes. In: Perlmann, G.E. and Lorand, L. (eds) Methods in Enzymology, Vol. XIX: Proteolytic Enzymes. Academic Press, New York, pp. 963–978.

236

T.S. George et al.

Demanèche, S., Jocteur-Monrozier, L., Quiquampoix, H. and Simonet, P. (2001) Evaluation of biological and physical protection against nuclease degradation of clay-bound plasmid DNA. Applied and Environmental Microbiology 67, 293–299. Dharmsthiti, S., Charlermpornpeisarn, S., Kiatiyajarn, M., Chanpokapaiboon, A., Klongsithidej, Y. and Techawiparut, J. (2005) Phytase production from Pseudomonas putida harbouring Escherichia coli appA. Process Biochemistry 40, 789–793. Dick, W.A. and Tabatabai, M.A. (1987) Kinetics and activities of phosphatase–clay complexes. Soil Science 143, 5–15. Dick, W.A., Juma, N.G. and Tabatabai, M.A. (1983) Effects of soils on acid phosphatase and inorganic pyrophosphatase of corn roots. Soil Science 136, 19–25. Dox, A.W. and Golden, R. (1911) Phytase in lower fungi. Journal of Biological Chemistry 10, 183–186. Eivazi, F. and Tabatabai, M.A. (1977) Phosphatases in soils. Soil Biology and Biochemistry 9, 167–172. Engasser, J.-M. and Horvath, C. (1976) Diffusion and kinetics with immobilised enzymes. In: Wingard, L.B. Jr, Katchalski-Katzir, E. and Goldstein, L. (eds) Applied Biochemistry and Bioengineering, Vol.1: Immobilized Enzyme Principles. Academic Press, New York, pp. 127–220. Erlich, K.C., Montalbano, B.G., Mullaney, E.J., Dischinger, H.C. and Ullah, A.H.J. (1993) Identification and cloning of a second phytase gene (phyB) from Aspergillus niger (ficuum). Biochemical and Biophysical Research Communications 195, 53–57. Espeland, E.M. and Wetzel, R.G. (2001) Complexation, stabilisation and UV photolysis of extracellular and surface bound glucosidase and alkaline phosphatase: implications for biofilm microbiota. Microbial Ecology 42, 572–585. Estermann, E.F., Peterson, G.H. and McLaren, A.D. (1959) Digestion of clay–protein, lignin–protein and silica–protein complexes by enzymes and bacteria. Soil Science Society of America Proceedings 23, 31–36. Feller, C., Frossard, E. and Brossard, M. (1994) Activite phosphatasique de quelques sols tropicaux à argile 1:1. Répartition dans les fractions granulométriques. Canadian Journal of Soil Science 74, 121–129. Findenegg, G.R. and Nelemans, J.A. (1993) The effect of phytase on the availability of phosphorus from myo-inositolhexaphosphate (phytate) in maize roots. Plant and Soil 154, 189–196. Freeman, C., Liska, G., Ostle, N.J., Lock, M.A., Reynolds, B. and Hudson, J. (1996) Microbial activity and enzymatic decomposition processes following peatland water table drawdown. Plant and Soil 180, 121–127. Geiger, G., Brandl, H., Furrer, G. and Schulin, R. (1998a) The effect of copper on the activity of cellulase and βglucosidase in the presence of montmorillonite or Al-montmorillonite. Soil Biology and Biochemistry 30, 1537–1544. Geiger, G., Livingston, M.P., Funk, F. and Schulin, R. (1998b) β-Glucosidase activity in the presence of copper and goethite. European Journal of Soil Science 49, 17–23. George, T.S., Richardson, A.E., Hadobas, P. and Simpson, R.J. (2004) Characterisation of transgenic Trifolium subterraneum L. which expresses phyA and releases extracellular phytase: growth and P nutrition in laboratory media and soil. Plant Cell and Environment 27, 1351–1361. George, T.S., Simpson, R.J., Hadobas, P.A. and Richardson, A.E. (2005a) Expression of a fungal phytase gene in Nicotiana tobacum improves phosphorus nutrition of plants grown in amended soils. Plant Biotechnology Journal 3, 129–140. George, T.S., Richardson, A.E. and Simpson, R.J. (2005b) Behaviour of plant-derived extracellular phytase upon addition to soil. Soil Biology Biochemistry 37, 977–988. George, T.S., Richardson, A.E., Smith, J.B., Hadobas, P.A. and Simpson, R.J. (2005c) Limitations to the potential of transgenic Trifolium subterraneum L. plants that exude phytase, when grown in soils with a range of organic P content. Plant and Soil 278, 263–274. George, T.S., Gregory, P.J., Simpson, R.J. and Richardson, A.E. (2006) Differential interaction of Aspergillus niger and Peniophora lycii phytases with soil particles affects the hydrolysis of inositol phosphates. Applied and Environmental Microbiology (in press). Gianfreda, L. and Bollag, J.-M. (1994) Effects of soil on the behaviour of immobilised enzymes. Soil Science Society of America Journal 58, 1672–1681. Gianfreda, L. and Bollag, J.-M. (1996) Influence of natural and anthropogenic factors on enzyme activity in soil. In: Stotsky, G. and Bollag, J.-M. (eds) Soil Biochemistry, Vol. 9. Marcel Dekker, New York, pp. 123–194. Gianfreda, L., Rao, M.A. and Violante, A. (1993) Interactions of invertase with tannic acid, hydroxy-aluminium (OH–Al) species or montmorillonite. Soil Biology and Biochemistry 25, 671–677. Gibbins, L.N. and Norris, F.W. (1963) Phytase and acid phosphatase in dwarf bean, Phaseolus vulgaris. Biochemical Journal 86, 67–71.


237

Gibson, D.M. and Ullah, A.H.J. (1988) Purification and characterization of phytase from cotyledons of germinating soybean seeds. Archives of Biochemistry and Biophysics 260, 503–513. Goel, R., Mino, T., Satoh, H. and Matsuo, T. (1998) Enzyme activities under anaerobic and aerobic conditions in activated sludge sequencing batch reactor. Water Research 32, 2081–2088. Goldstein, L. (1976) Kinetic behaviour of immobilised enzyme systems. In: Mosbach, K. (ed.) Methods in Enzymology, Vol. XLIV: Immobilized Enzymes. Academic Press, New York, pp. 397–443. Golovan, S.P., Hayes, M.A., Phillips, J.P. and Forsberg, C.W. (2001) Transgenic mice expressing bacterial phytase as a model for phosphorus pollution control. Nature Biotechnology 19, 429–432. Greiner, R., Konietzny, U. and Jany, K.-D. (1993) Purification and characterisation of two phytases from Escherichia coli. Archives of Biochemistry and Biophysics 303, 107–113. Han, Y., Wilson, D.B. and Lei, X.G. (1999) Expression of an Aspergillus niger phytase gene (phyA) in Saccharomyces cerevisiae. Applied and Environmental Microbiology 65, 1915–1918. Harrison, A.F. (1983) Relationship between intensity of phosphatase activity and physico-chemical properties in woodland soils. Soil Biology and Biochemistry 15, 93–99. Harrison, A.F. (1987) Soil Organic Phosphorus: A Review of World Literature. CAB International, Wallingford, UK. Hayes, J.E., Richardson, A.E. and Simpson, R.J. (1999) Phytase and acid phosphatase activities in extracts from roots of temperate pasture grass and legume seedlings. Australian Journal of Plant Physiology 26, 801–809. Huang, Q. and Shindo, H. (2000a) Effects of copper on the activity and kinetics of free and immobilised acid phosphatase. Soil Biology and Biochemistry 32, 1885–1892. Huang, Q. and Shindo, H. (2000b) Inhibition of free and immobilised acid phosphatase by zinc. Soil Science 165, 793–802. Huang, Q. and Shindo, H. (2001) Comparison of the influence of Cu, Zn, and Cd on the activity and kinetics of free and immobilised acid phosphatase. Soil Science and Plant Nutrition (Tokyo) 47, 767–772. Huang, Q., Jiang, M. and Li, X. (1999) Adsorption and properties of urease immobilised on several iron and aluminium oxides (hydroxides) and kaolinite. In: Berthelin, J., Huang, P.M., Bollag, J.-M. and Andreux, F. (eds) Effects of Mineral–Organic–Microorganism Interactions on Soil and Freshwater Environments. Kluwer Academic Press, Dordrecht, The Netherlands, pp. 167–174. Huang, Q., Chen, W., Gianfreda, L. and Violante, A. (2002) Adsorption of acid phosphatase on minerals and soil colloids in presence of citrate and phosphate. Pedosphere 12, 339–348. Huang, Q., Zhao, Z. and Chen, W. (2003) Effects of several low-molecular weight organic acids and phosphate on the adsorption of acid phosphatase by soil colloids and minerals. Chemosphere 52, 571–579. Idriss, E.E., Makarewicz, O., Farouk, A., Rosner, K., Greiner, R., Bochow, H., Richter, T. and Borriss, R. (2002) Extracellular phytase activity of Bacillus amyloloquefaciens FZB45 contributes to its plant-growth-promoting effect. Microbiology 148, 2097–2109. Irving, G.C.J. (1980) Phytase. In: Cosgrove, D.J. (ed.) Inositol Phosphates – Their Chemistry, Biochemistry, and Physiology. Elsevier, Amsterdam, The Netherlands, pp. 85–117. Irving, G.C.J. and Cosgrove, D.J. (1971a) Inositol phosphate phosphatases of microbiological origin: some properties of a partially purified bacterial (Pseudomonas sp.) phytase. Australian Journal of Biological Science 24, 547–557. Irving, G.C.J. and Cosgrove, D.J. (1971b) Inositol phosphate phosphatases of microbiological origin: observations on the nature of the active centre of a bacterial (Pseudomonas sp.) phytase. Australian Journal of Biological Science 24, 559–564. Irving, G.C.J. and Cosgrove, D.J. (1974) Inositol phosphate phosphatases of microbiological origin: some properties of the partially purified phosphatases of Aspergillus ficuum NRRL 3135. Australian Journal of Biological Science 27, 361–368. Jackman, R.H. and Black, C.A. (1952) Phytase activity in soils. Soil Science 73, 117–125. Juma, N.G. and Tabatabai, M.A. (1977) Effects of trace elements on phosphatase activity in soils. Soil Science Society of America Journal 41, 343–349. Kanazawa, S. and Filip, Z. (1986) Distribution of microorganisms, total biomass and enzyme activities in different particles of brown soil. Microbial Ecology 12, 205–215. Kandeler, E. (1990) Characterization of free and adsorbed phosphatases in soils. Biology and Fertility of Soils 9, 199–202. Kang, H. and Freeman, C. (1999) Phosphatase and arylsulphatase activities in wetland soils: annual variation and controlling factors. Soil Biology and Biochemistry 31, 449–454. Kelleher, B.P., Simpson, A.J., Willeford, K.O., Simpson, M.J., Stout, R., Rafferty, A. and Kingery, W.L. (2004) Acid phosphatase interactions with organo-mineral complexes: influence on catalytic activity. Biogeochemistry 71, 285–297.

238

T.S. George et al.

Kerovuo, J., Lauraeus, M., Nurminen, P., Kalkkinen, N. and Apajalahti, J. (1998) Isolation, characterisation, molecular gene cloning and sequencing of a novel phytase from Bacillus subtilis. Applied and Environmental Microbiology 64, 2079–2085. Kerovuo, J., Lappalainen, I. and Reinikainen, T. (2000) The metal dependence of Bacillus subtilis phytase. Biochemical and Biophysical Research Communications 268, 365–369. Kim, Y.O., Kim, H.K., Bae, K.S., Yu, J.H. and Oh, T.K. (1998a) Purification and properties of a thermostable phytase from Bacillus sp. DS11. Enzyme and Microbial Technology 22, 2–7. Kim, Y.O., Lee, J.K., Kim, H.K., Yu, J.H. and Oh, T.K. (1998b) Cloning of the thermostable phytase gene (phy) from Bacillus sp. DS11 and its overexpression in Escherichia coli. FEMS Microbiology Letters 162, 185–191. Kim, Y.O., Lee, J.K., Oh, B.C. and Oh, T.K. (1999) High-level expression of a recombinant thermostable phytase in Bacillus subtilis. Bioscience, Biotechnology, and Biochemistry 63, 2205–2207. Kondo, A. and Higashitani, K. (1992) Adsorption of model proteins with wide variation in molecular properties on colloidal particles. Journal of Colloid and Interface Science 150, 344–351. Kondo, A., Murakami, F., Kawagoe, M. and Higashitani, K. (1993) Kinetic and circular dichroism studies of enzymes adsorbed on ultrafine silica particles. Applied Microbiological Biotechnology 39, 726–731. Ladd, J.N. and Butler, J.H.A. (1975) Humus-enzyme systems and synthetic, organic polymer–enzyme analogs. In: Paul, E.A. and McLaren, A.D. (eds) Soil Biochemistry, Vol. 4. Marcel Dekker, New York, pp. 143–194. Ladd, J.N., Foster, R.C. and Skjemstad, J.O. (1993) Soil structure: carbon and nitrogen metabolism. Geoderma 56, 401–434. Ladd, J.N., Foster, R.C., Nannipieri, P. and Oades, J.M. (1996) Soil structure and biological activity. In: Stotzky, G. and Bollag, J.-M. (eds) Soil Biochemistry, Vol. 9. Marcel Dekker, New York, pp. 23–78. Laidler, J. and Sundaram, P.V. (1971) The kinetics of supported enzyme systems. In: Brown, H.D. (ed.) Chemistry of the Cell Interface. Academic Press, New York, pp. 255–296. Lassen, S.F., Breinholt, J., Østergaard, P.R., Brugger, R., Bischoff, A., Wyss, M. and Fuglsang, C.C. (2001) Expression, gene cloning, and characterization of five novel phytases from four basidiomycete fungi: Peniophora lycii, Agrocybe pediades, a Ceriporia sp., and Trametes pubescens. Applied and Environmental Microbiology 67, 4701–4707. Lehmann, M., Pasamontes, L., Lassen, S.F. and Wyss, M. (2000) The consensus concept for thermostability engineering of proteins. Biochimica et Biophysica Acta 1543, 408–415. Lehmann, M., Loch, C., Middendorf, A., Studer, D., Lassen, S.F., Pasamontes, L., van Loon, A.P.G.M. and Wyss, M. (2002) The consensus concept for thermostability engineering of proteins: further proof of concept. Protein Engineering 15, 403–411. Lei, X.G. and Porres, J.M. (2003) Phytase enzymology, applications and biotechnology. Biotechnology Letters 25, 1787–1794. Leprince, F. and Quiquampoix, H. (1996) Extracellular enzyme activity in soil: effect of pH and ionic strength on the interaction with montmorillonite of two acid phosphatases secreted by the ectomycorrhizal fungus Hebeloma cylindrosporum. European Journal of Soil Science 47, 511–522. Liu, B.-L., Jong, C.-H. and Tzeng, Y.M. (1999) Effect of immobilisation on pH and thermal stability of Aspergillus ficuum phytase. Enzyme and Microbial Technology 25, 517–521. Lolas, G.M. and Markakis, P. (1977) The phytase of navy beans (Phaseolus vulgaris). Journal of Food Science 42, 1094–1097. Lung, S.-C., Chan, W.-L., Yip, W., Wang, L., Yeung, E.C. and Lim, B.L. (2005) Secretion of beta-propeller phytase from tobacco and Arabidopsis roots enhances phosphorus utilisation. Plant Science 169, 341–349. Makboul, H.E. and Ottow, J.C.G. (1979) Alkaline phosphatase activity and Michaelis constant in the presence of different clay minerals. Soil Science 128, 129–135. Mandal, N.C., Burman, S. and Biswas, B.B. (1972) Isolation, purification and characterisation of phytase from germinating mung beans. Phytochemistry 11, 495–502. Marx, M.-C., Kandeler, E., Wood, M., Wermbter, N. and Jarvis, S.C. (2005) Exploring the enzymatic landscape: distribution and kinetics of hydrolytic enzymes in soil particle-size fractions. Soil Biology and Biochemistry 37, 35–48. Marzadori, C., Gessa, C. and Ciurli, S. (1998) Kinetic properties and stability of potato acid phosphatase immobilised on Ca-polygalacturonate. Biology and Fertility of Soils 27, 97–103. Matumoto-Pintro, P.T. and Quiquampoix, H. (1997) La phase solide des sols comme contrainte au fonctionnement des enzymes sécrétés par les microorganismes: comparaison de phytases intra et extracellulaires. In: Baleux, B., Desmazeaud, M., Divies, C., Gendre, F. and Moletta, R. (eds) Microbiologie Industrielle et Environnement. Sociétét Française de Microbiologie, Paris, France, pp. 195–204.


239

McLaren, A.D. and Packer, L. (1970) Some aspects of enzyme reaction in heterogenous systems. Advances in Enzymology 33, 245–308. Mullaney, E.J., Daly, C.B., Kim, J.M., Porres, J.M., Lei, X.G., Sethumadhavan, K. and Ullah, A.H.J. (2002) Sitedirected mutagenesis of Aspergillus niger NRRL 3135 phytase at residue 300 to enhance catalysis at pH 4.0. Biochemical and Biophysical Research Communications 297, 1016–1020. Nagai, Y. and Funahashi, S. (1962) Phytase from wheat bran. I. Purification and substrate specificity. Agricultural and Biological Chemistry 26, 794–803. Naidja, A., Huang, P.M. and Bollag, J.M. (2000) Enzyme–clay interactions and their impact on transformations of natural and anthropogenic organic compounds in soil. Journal of Environmental Quality 29, 677–691. Nannipieri, P. and Gianfreda, L. (1998) Kinetics of enzyme reactions in soil environments. In: Huang, P.M., Senesi, N. and Buffle, J. (eds) Structure and Surface Reactions of Soil Particles. John Wiley & Sons, Chichester, UK, pp. 449–479. Nannipieri, P., Ceccanti, B. and Bianchi, D. (1988) Characterisation of humus–phosphatase complexes extracted from soil. Soil Biology and Biochemistry 20, 663–691. Nannipieri, P., Sequi, P. and Fusi, P. (1996) Humus and enzyme activity. In: Piccolo, A. (ed.) Humic Substances in Terrestrial Ecosystems. Elsevier, Amsterdam, The Netherlands, pp. 293–328. Nayini, N.R. and Markakis, P. (1984) The phytase of yeast. Lebensmittel Wissenschaft und Technologie 17, 24–26. Nayini, N.R. and Markakis, P. (1986) Phytases. In: Graf, E. (ed.) Phytic Acid: Chemistry and Applications. Pilatus Press, Minneapolis, Minnesota, pp. 101–118. Noinville, S., Revault, M., Quiquampoix, H. and Baron, M.H. (2004) Structural effects of drying and rehydration for enzymes in soils: a kinetics-FTIR analysis of α-chymotrypsin adsorbed on montmorillonite. Journal of Colloid and Interface Science 273, 414–425. Norde, W. and Lyklema, J. (1991) Why proteins prefer interfaces? Journal of Biomaterial Science, Polymer Edition 2, 183–202. Olander, L.P. and Vitousek, P.M. (2000) Regulation of soil phosphatase and chitinase activity by N and P availability. Biogeochemistry 49, 175–190. Peers, F.G. (1953) The phytase of wheat. The Biochemical Journal 53, 102–110. Pen, J., Verwoerd, T.C., van Paridon, P.A., Beudeker, R.F., van den Elzen, P.J.M., Geerse, K., van der Klis, J.D., Versteeghm, H.A.J., van Ooyen, A.J.J. and Hoekema, A. (1993) Phytase-containing transgenic seeds as a novel feed additive for improved phosphorus utilization. BiolTechnology 11, 811–814. Perez-Mateos, M. and Rad, J.C. (1989) Immobilisation of alkaline phosphatase by soil structural units. Biotechnology and Applied Biochemistry 11, 371–378. Perez-Mateos, M., Busto, M. and Rad, J.C. (1991) Stability and properties of alkaline phosphatase immobilised on a rendzina soil. Journal of the Science of Food and Agriculture 55, 229–240. Pettit, N.M., Gregory, L.J., Freedman, R.B. and Burns, R.G. (1977) Differential stabilities of soil enzymes: assay and properties of phosphatase and arylsulphatase. Biochimica et Biophysica Acta 485, 357–366. Powar, V.K. and Jagannathan, V. (1982) Purification and properties of phytate-specific phosphatase from Bacillus subtilis. Journal of Bacteriology 151, 1102–1108. Pulford, I.D. and Tabatabai, M.A. (1988) Effect of waterlogging on enzyme activities in soils. Soil Biology and Biochemistry 20, 215–219. Quan, C.-S., Tian, W.-J., Fan, S.-D. and Kikuchi, J.-I. (2004) Purification and properties of a low-molecularweight phytase from Cladosporium sp. FP-1. Journal of Bioscience and Bioengineering 97, 260–266. Quiquampoix, H. (1987a) A stepwise approach to the understanding of extracellular enzyme activity in soil. I. Effect of electrostatic interactions on the conformation of a β-D-glucosidase on different mineral surfaces. Biochimie 69, 753–763. Quiquampoix, H. (1987b) A stepwise approach to the understanding of extracellular enzyme activity in soil. II. Competitive effects of the adsorption of a β-D-glucosidase in mixed mineral or organo-mineral systems. Biochimie 69, 765–771. Quiquampoix, H. (2000) Mechanisms of protein adsorption on surfaces and consequences for extracellular enzyme activity in soil. In: Bollag, J.-M. and Stotzky, G. (eds) Soil Biochemistry, Vol. 10. Marcel Dekker, New York, pp. 171–206. Quiquampoix, H. and Mousain, D. (2005) Enzymatic hydrolysis of organic phosphorus. In: Turner, B.L., Frossard, E. and Baldwin, D. (eds) Organic Phosphorus in the Environment. CAB International, Wallingford, UK, pp. 89–112. Quiquampoix, H., Staunton, S., Baron, M.-H. and Ratcliffe, R.G. (1993) Interpretation of the pH dependence of protein adsorption on clay mineral surfaces and its relevance to the understanding of extracellular enzyme activity in soil. Colloids and Surfaces A: Physicochemical and Engineering Aspects 75, 85–93.

240

T.S. George et al.

Quiquampoix, H., Abadie, J., Baron, M.-H., Leprince, F., Matumoto-Pintro, P.T., Ratcliffe, R.G. and Staunton, S. (1995) Mechanisms and consequences of protein adsorption on soil mineral surfaces. American Chemical Society Symposium Series 602, 321–333. Quiquampoix, H., Servagent-Noinville, S. and Baron, M.-H. (2002) Enzyme adsorption on soil mineral surfaces and consequences for the catalytic activity. In: Burns, R.G. and Dick, R.P. (eds) Enzymes in the Environment: Activity, Ecology and Applications. Marcel Dekker, New York, pp. 285–306. Rao, M.A. and Gianfreda, L. (2000) Properties of acid phosphatase–tannic acid complexes formed in the presence of Fe and Mn. Soil Biology and Biochemistry 32, 1921–1926. Rao, M.A., Violante, A. and Gianfreda, L. (1994) Catalytic behaviour of acid phosphatase immobilized on clay minerals and organo-mineral complexes. Transactions of the 15th World Congress of Soil Science. Acapulco, Mexico, 10–16 July 1994, Vol. 3b. International Soil Science Society, Mexico, pp. 117–118. Rao, M.A., Violante, A. and Gianfreda, L. (2000) Interaction of acid phosphatase with clays, organic molecules and organo-mineral complexes: kinetics and stability. Soil Biology and Biochemistry 32, 1007–1014. Rao, M.A., Sannino, F., Nocerino, E., Puglisi, E. and Gianfreda, L. (2003) Effect of air-drying treatment on enzymatic activities of soils affected by anthropogenic activities. Biology and Fertility of Soils 38, 327–332. Renella, G., Ortigoza, A.L.R., Landi, L. and Nannipieri, P. (2003) Additive effects of copper and zinc on cadmium toxicity on phosphatase activities and ATP content of soil as estimated by the ecological dose (ED50). Soil Biology and Biochemistry 35, 1203–1210. Reyes, J.L., Rodrigo, M.-J., Colmenero-Flores, J.M., Gil, J.-V., Garay-Arroyo, A., Campos, F., Salamini, F., Bartels, D. and Covarrubias, A.A. (2005) Hydrophillins from distant organisms can protect enzymatic activities from water limitation effects in vitro. Plant Cell and Environment 28, 709–718. Richardson, A.E., Hadobas, P.A. and Hayes, J.E. (2001a) Extracellular secretion of Aspergillus phytase from Arabidopsis roots enables plants to obtain phosphorus from phytate. The Plant Journal 25, 641–649. Richardson, A.E., Hadobas, P.A., Hayes, J.E., O’Hara, C.P. and Simpson, R.J. (2001b) Utilization of phosphorus by pasture plants supplied with myo-inositol hexaphosphate is enhanced by the presence of soil microorganisms. Plant Soil 229, 47–56. Rojo, M.J., Carcedo, S.G. and Mateos, M.P. (1990) Distribution and characterisation of phosphatase and organic phosphorus in soil fractions. Soil Biology and Biochemistry 22, 169–174. Rowell, D.L. (1994) Soil Science: Methods and Applications. Longman Group, Marlow, UK. Saa, A., Trasar-Cepeda, C., Gil-Sotres, F. and Carballas, T. (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. 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. Staddon, W.J., Duchesne, L.C. and Trevors, J.T. (1998) Acid phosphatase, alkaline phosphatase and arylsulfatase activities in soils from jack pine (Pinus banksiana Lamb.) ecosystem after clearing, prescribed burning and scarification. Biology and Fertility of Soils 27, 1–4. Suzuki, U., Yoshimura, K. and Takaishi, M. (1907) Ueber ein enzym ‘Phytase’ das ‘Anhydoro-oxy-methylen diphosphorsaure’ spaltet. Bulletins of the College of Agriculture Tokyo 5, 503–512. Svenson, A. (1986) Effects of copper, zinc and cadmium ions on the production of phosphate from phytic acid by the phytase system in spruce forest soil. Plant and Soil 94, 227–234. Tarafdar, J.C., Yadav, R.S. and Niwas, R. (2002) Relative efficiency of fungal intra- and extracellular phosphatases and phytase. Journal of Plant Nutrition and Soil Science 165, 17–19. Tietjen, T. and Wetzel, R.G. (2003) Extracellular enzyme–clay mineral complexes: enzyme adsorption, alteration of enzyme activity and protection from photodegredation. Aquatic Ecology 37, 331–339. Tinker, P.B. and Nye, P.H. (2000) Solute Movement in the Rhizosphere. Oxford University Press, Oxford.


241

Tomschy, A., Brugger, R., Lehmann, M., Svendsen, A., Vogel, K., Kostrewa, D., Lassen, S.F., 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. Ullah, A.J.H. and Cummins, B.J. (1987) Purification, N-terminal amino acid sequence and characterization of pH 2.5 optimum acid phosphatase (E.C. 3.1.3.2) from Aspergillus ficuum. Preparative Biochemistry 17, 397–422. Ullah, A.J.H. 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. Vats, P. and Banerjee, U.C. (2004) Production studies and catalytic properties of phytases (myo-inositol hexakisphosphate phosphatases) an overview. Enzyme and Microbial Technology 35, 3–14. Violante, A. and Gianfreda, L. (2000) Role of biomolecules in the formation and reactivity toward nutrients and organics of variable charge minerals and organo mineral complexes in soil environments. In: Bollag, J.-M. and Stotsky, G. (eds) Soil Biochemistry, Vol. 10. Marcel Dekker, New York, pp. 107–270. Violante, A., De Cristofaro, A., Rao, M.A. and Gianfreda, L. (1995) Physicochemical properties of protein–smectite and protein–Al(OH)x–smectite complexes. Clay Minerals 30, 325–336. Wetzel, R.G. (1992) Gradient-dominated ecosystems: sources and regulatory functions of dissolved organic matter in freshwater ecosystems. Hydrobiologia 229, 181–198. Wetzel, R.G. (1993) Humic compounds from wetlands: complexation, inactivation and reactivation of surfacebound and extracellular enzymes. Verhandlungen der Internationale Vereinigung für Liminologie 25, 122–128. Wyss, M., Pasamontes, L., Remy, R., Kohler, J., Kusznir, E., Gadient, M., Muller, F. and van Loon, A.P.G.M. (1998) Comparisons of the thermostability properties of three acid phosphatases from molds: Aspergillus fumigatus phytase, A. niger phytase and A. niger pH 2.5 acid phosphatase. Applied and Environmental Microbiology 64, 4446–4451. Wyss, M., Brugger, R., Kronenberger, A., Remy, R., Fimbel, R., Gottfried, O., Lehmann, M. and van Loon, A.P.G.M. (1999a) Biochemical characterisation of fungal phytases (myo-inositol hexakisphosphate phosphatases): 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., Schnobelen, 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 phosphatases): molecular size, glycosylation pattern and engineering of proteolytic resistance. Applied and Environmental Microbiology 65, 359–366. Yamada, K., Minoda, Y. and Yamamoto, S. (1968) Phytase from Aspergillus terreus. Part I. Production, purification and some general properties of the enzyme. Agricultural and Biological Chemistry 32, 1275–1282. 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. Zimmermann, P., Zardi, G., Lehmann, M., Zelder, 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.

15

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



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


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).

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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


myo-Inositol hexakisphosphate (sodium-phytate)

No P

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ex::phyA

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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


249

Calciumphytate

Nicotiana tabacum

Alfisol phosphate

No P

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250

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Calcium-phytate 200 Alfisol 100

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125

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Calcium-phytate 200 Spodosol

100

0 No P

15.5

62.5

125

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,


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

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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


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.

References Adams, M.A. and Pate, J.S. (1992) Availability of organic and inorganic forms of phosphorus to lupins (Lupinus spp.). Plant and Soil 145, 107–113. Alvarez, M., Godoy, R., Heyser, W. and Härtel, S. (2005) Anatomical–physiological determination of surface bound phosphatase activity in ectomycorrhizae of Nothofagus obliqua. Soil Biology and Biochemistry 37, 125–132. 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. 411–432. Anderson, G., Williams, E.G. and Moir, J.O. (1974) A comparison of the sorption of inorganic orthophosphate and inositol hexaphosphate by six acid soils. Journal of Soil Science 25, 51–62. Antibus, R.K., Croxdale, J.G., Miller, O.K. and Linkins, A.E. (1981) Ectomycorrhizal fungi of Salix rotundifolia. III. Resynthesized mycorrhizal complexes and their surface phosphatase activities. Canadian Journal of Botany 59, 2458–2465. Antibus, R.K., Sinsabaugh, R.L. and Linkins, A.E. (1992) Phosphatase activities and phosphorus uptake from inositol phosphates by ectomycorrhizal fungi. Canadian Journal of Botany 70, 794–801. Antibus, R.K., Bower, D. and Dighton, J. (1997) Root surface phosphatase activities and uptake of 32P-labelled inositol phosphate in field-collected gray birch and red maple roots. Mycorrhiza 7, 39–46. Asmar, F. (1997) Variation in activity of root extracellular phytase between genotypes of barley. Plant and Soil 195, 61–64. Barrett-Lennard, E.G., Dracup, M. and Greenway, H. (1993) Role of extracellular phosphatase in the phosphorusnutrition of clover. Journal of Experimental Botany 44, 1595–1600. Bartlett, E.M. and Lewis, D.H. (1973) Surface phosphatase activity of mycorrhizal roots of beech. Soil Biology and Biochemistry 5, 249–257. Baxter, J.W. and Dighton, J. (2005) Phosphorus source alters host plant response to ectomycorrhizal diversity. Mycorrhiza 15, 513–523.


255

Beck, E., Fußeder, A. and Kraus, M. (1989) The maize root system in situ: evaluation of structure and capability of utilization of phytate and inorganic phosphates. Zeitschrift für Pflanzenernährung und Bodenkunde 152, 159–167. Bieleski, R.L. (1973) Phosphate pools, phosphate transport and phosphate availability. Annual Reviews of Plant Physiology 24, 225–252. Bolan, N.S. (1991) A critical review on the role of mycorrhizal fungi in the uptake of phosphorus by plants. Plant and Soil 134, 189–207. Bosse, D. and Köck, M. (1998) Influence of phosphate starvation on phosphohydrolases during development of tomato seedlings. Plant Cell and Environment 21, 325–332. Campbell, M., Dunn, R., Ditterline, R., Pickett, S. and Raboy, V. (1991) Phytic acid represents 10% to 15% of total phosphorus in alfalfa root and crown. Journal of Plant Nutrition 14, 925–937. Celi, L. and Barberis, E. (2005) Abiotic stabilization of organic phosphorus in the environment. In: Turner, B.L., Frossard, E. and Baldwin, D.S. (eds) Organic Phosphorus in the Environment. CAB International, Wallingford, UK, pp. 113–132. Chen, A., Chambers, S.M., and Cairney, J.W.G. (1999) Utilisation of organic nitrogen and phosphorus sources by mycorrhizal endophytes of Woollsia pungens (Cav.) F. Muell. (Epacridaceae). Mycorrhiza 8, 181–187. Chen, C.R., Condron, L.M., Davis, M.R. and Sherlock, R.R. (2002) Phosphorus dynamics in the rhizosphere of perennial ryegrass (Lolium perenne L.) and radiata pine (Pinus radiata D. Don). Soil Biology and Biochemistry 34, 487–499. Chen, C.R., Condron, L.M., Turner, B.L., Mahieu, N., Davis, M.R., Xu, Z.H. and Sherlock, R.R. (2004) Mineralisation of soil orthophosphate monoesters under pine seedlings and ryegrass. Australian Journal of Soil Research 42, 189–196. Colpaert, J.V., van Laere, A., van Tichelen, K.K. and van Assche, J.A. (1997) The use of inositol hexaphosphate as a phosphorus source by mycorrhizal and non-mycorrhizal Scots pine (Pinus sylvestris). Functional Ecology 11, 407–415. Cosgrove, D. J., Irving, G.C.J. and Bromfield, S.M. (1970) Inositol phosphate phosphatases of microbial origin: the isolation of soil bacteria having inositol phosphate phosphatase activity. Australian Journal of Biological Sciences 23, 339–343. Dinkelaker, B. and Marschner, H. (1992) In vivo demonstration of acid phosphatase activity in the rhizosphere of soil-grown plants. Plant and Soil 144, 199–205. Dodd, J.C., Burton, C.C., Burns, R.G. and Jeffries, P. (1987) Phosphatase activity associated with the roots and the rhizosphere of plants infected with vesicular arbuscular mycorrhizal fungi. New Phytologist 107, 163–172. Dracup, M.N.H., Barrett-Lennard, E.G., Greenway, H. and Robson, A.D. (1984) Effect of phosphorus deficiency on phosphatase activity of cell walls from roots of subterranean clover. Journal of Experimental Botany 35, 466–480. Espinosa, M., Turner, B.L. and Haygarth, P.M. (1999) Preconcentration and separation of trace phosphorus compounds in soil leachate. Journal of Environmental Quality 28, 1497–1504. Ezawa, T. and Yoshida, T. (1994) Acid phosphatase specific to arbuscular mycorrhizal infection in marigold and possible role in symbiosis. Soil Science and Plant Nutrition 40, 655–665. Ezawa, T., Hayatsu, M. and Saito, M. (2005) A new hypothesis on the strategy for acquisition of phosphorus in arbuscular mycorrhiza: up-regulation of secreted acid phosphatase gene in the host plant. Molecular Plant–microbe Interactions 18, 1046–1053. Feng, G., Su, Y.B., Li, X.L., Wang, H., Zhang, F.S., Tang, C. and Rengel, Z. (2002) Histochemical visualization of phosphatase released by arbuscular mycorrhizal fungi in soil. Journal of Plant Nutrition 25, 969–980. Feng, G., Song, Y.C., Li, X.L. and Christie, P. (2003) Contribution of arbuscular mycorrhizal fungi to utilization of organic sources of phosphorus by red clover in calcareous soil. Applied Soil Ecology 22, 139–148. Findenegg, G.R. and Nelemans, J.A. (1993) The effect of phytase on the availability of phosphorus from myo-inositol hexaphosphate (phytate) for maize roots. Plant and Soil 154, 189–196. Firsching, B.M. and Claassen, N. (1996) Root phosphatase activity and soil organic phosphorus utilization by Norway spruce (Picea abies (L.) Karst.). Soil Biology and Biochemistry 28, 1417–1424. Fries, L.L.M., Pacovsky, R.S., Safir, G.R. and Kaminski, J. (1998) Phosphorus effect on phosphatase activity in endomycorrhizal maize. Physiologia Plantarum 103, 162–71. Gavito, M.E. and Olsson, P.A. (2003) Allocation of plant carbon to foraging and storage in arbuscular mycorrhizal fungi. FEMS Microbiology Ecology 45, 181–187. George, T.S., Gregory, P. J., Robinson, J.S. and Buresh, R. J. (2002) Changes in phosphorus concentrations and pH in the rhizosphere of some agroforestry and crop species. Plant and Soil 246, 65–73.

256


George, T.S., Richardson, A.E., Hadobas, P.A. and Simpson, R. J. (2004) Characterisation of transgenic Trifolium subterraneum L. which expresses phyA and releases extracellular phytase: growth and phosphorus nutrition in laboratory media and soil. Plant, Cell and Environment 27, 1351–1361. George, T.S., Richardson, A.E. and Simpson, R. J. (2005a) Behaviour of plant-derived extracellular phytase upon addition to soil. Soil Biology and Biochemistry 37, 977–988. George, T.S., Richardson. A.E., Smith, J.B., Hadobas, P.A. and Simpson, R. J. (2005b) Limitations to the potential of transgenic Trifolium subterraneum L. plants that exude phytase, when grown in soils with a range of organic phosphorus content. Plant and Soil 278, 263–274. George, T.S., Simpson, R. J., Hadobas, P.A. and Richardson, A.E. (2005c) Expression of a fungal phytase gene in Nicotiana tobacum improves phosphorus nutrition in plants grown in amended soil. Plant Biotechnology Journal 3, 129–140. George, T.S., Turner, B.L., Gregory, P. and Richardson, A.E. (2006a) Depletion of organic phosphorus from oxisols in relation to phosphatase activities in the rhizosphere. European Journal of Soil Science 57, 47–57. George, T.S., Gregory, P. J., Simpson, R. J. and Richardson, A.E. (2006b) Differential interactions of Aspergillus niger and Peniophora lycii with soil particles affects the hydrolysis of inositol phosphates. Applied and Environmental Microbiology (in review). Gilbert, G.A., Knight, J.D., Vance, C.P. and Allan, D.L. (1999) Acid phosphatase activity in phosphorus-deficient white lupin roots. Plant, Cell and Environment 22, 801–810. Greaves, M.P. and Webley, D.M. (1969) The hydrolysis of myo-inositol hexaphosphate by soil microorganisms. Soil Biology and Biochemistry 1, 37–43. Grierson, P.F. and Comerford, N.B. (2000) Non-destructive measurement of acid phosphatase activity in the rhizosphere using nitrocellulose membranes and image analysis. Plant and Soil 218, 49–57. Haran, S., Logendra, S., Seskar, M., Bratanova, M. and Raskin, I. (2000) Characterization of Arabidopsis acid phosphatase promoter and regulation of acid phosphatase expression. Plant Physiology 124, 615–626. Harrison, A.F. (1987) Soil Organic Phosphorus: A Review of World Literature. CAB International, Wallingford, UK. Häussling, M. and Marschner, H. (1989) Organic and inorganic soil phosphates and acid phosphatases activity in the rhizosphere of 80-year-old Norway spruce (Picea abies (L.) Karst.) trees. Biology and Fertility of Soils 8, 128–133. Hayes, J.E., Richardson, A.E., and Simpson, R. J. (1999) Phytase and acid phosphatase activities in extracts from roots of temperate pasture grass and legume seedlings. Australian Journal of Plant Physiology 26, 801–809. Hayes, J.E., Richardson, A.E. and Simpson, R. J. (2000a) Components of organic phosphorus in soil extracts that are hydrolysed by phytase and acid phosphatase. Biology and Fertility of Soils 32, 279–286. Hayes, J.E., Simpson, R. J. and Richardson, A.E. (2000b) The growth and utilisation of plants in sterile media when supplied with inositol hexaphosphate, glucose 1-phosphate or inorganic phosphate. Plant and Soil 220, 165–174. Hegeman, C.E. and Grabau, E.A. (2001) A novel phytase with sequence similarity to purple acid phosphatases is expressed in cotyledons of germinating soybean seedlings. Plant Physiology 126, 1598–1608. Helal, H.M. and Dressler, A. (1989) Mobilization and turnover of soil phosphorus in the rhizosphere. Zeitschrift für Pflanzenernahrung und Bodenkunde 152, 175–180. Hens, M. and Merckx, R. (2001) Functional characterization of colloidal phosphorus species in the soil solution of sandy soils. Environmental Science and Technology 35, 493–500. Hill, J.O., Simpson, R. J., Moore, A.D. and Chapman, D.F. (2006) Morphology and response of roots of pasture species to phosphorus and nitrogen nutrition. Plant and Soil (in press). Hinsinger, P. (2001) Bioavailability of soil inorganic phosphorus in the rhizosphere as affected by root-induced chemical changes: a review. Plant and Soil 237, 173–195. Ho, I. (1989) Acid phosphatase, alkaline phosphatase, and nitrate reductase activity of selected ectomycorrhizal fungi. Canadian Journal of Botany 67, 750–753. Hocking, P. J. (2001) Organic acids exuded from roots in phosphorus uptake and aluminium tolerance of plants in acid soils. Advances in Agronomy 74, 63–97. Hodge, A. (2004) The plastic plant: root responses to heterogeneous supplies of nutrients. New Phytologist 162, 9–24. Hübel, F. and Beck, E. (1993) In-situ determination of the phosphorus-relations around the primary root of maize with respect to inorganic and phytate-phosphorus. Plant and Soil 157, 1–9. Hübel, F. and Beck, E. (1996) Maize root phytase: purification, characterization and localization of enzyme activity and its putative substrate. Plant Physiology 112, 1429–1436. Idriss, E.E., Makarewicz, O., Farouk, A., Rosner, K., Greiner, R., Bochow, H., Richter, T. and Borriss, R. (2002) Extracellular phytase activity of Bacillus amyloliquefaciens FZB45 contributes to its plant-growth-promoting effect. Microbiology 148, 2097–2109.


257

Jackman, R.H. and Black, C.A. (1951) Solubility of iron, aluminium, calcium and magnesium inositol phosphates at different pH values. Soil Science 72, 179–186. Jakobsen, I., Leggett, M.E. and Richardson, A.E. (2005) Rhizosphere microorganisms and plant phosphorus uptake. In: Sims, J.T. and Sharpley, A.N (eds) Phosphorus, Agriculture and the Environment. American Society for Agronomy, Madison, Wisconsin, pp. 437–494. Jayachandran, K., Schwab, A.P. and Hetrick, B.A.D. (1992) Mineralization of organic phosphorus by vesicular–arbuscular mycorrhizal fungi. Soil Biology and Biochemistry 24, 897–903. Joner, E. J. and Jakobsen, I. (1994) Contribution by two arbuscular mycorrhizal fungi to phosphorus uptake by cucumber (Cucumis sativus L) from P32-labelled organic matter during mineralization in soil. Plant and Soil 163, 203–209. Joner, E. J. and Jakobsen, I. (1995) Uptake of 32P from labeled organic matter by mycorrhizal and non-mycorrhizal subterranean clover (Trifolium subterraneum L). Plant and Soil 172, 221–227. Joner, E. J. and Johansen, A. (2000) Phosphatase activity of external hyphae of two arbuscular mycorrhizal fungi. Mycological Research 104, 81–86. Joner, E. J., Magid, J., Gahoonia, T.S. and Jakobsen, I. (1995) Phosphorus depletion and activity of phosphatases in the rhizosphere of mycorrhizal and non-mycorrhizal cucumber (Cucumis sativus L). Soil Biology and Biochemisty 27, 1145–1151. Joner, E. J., van Aarle, I.M. and Vosatka, M. (2000) Phosphatase activity of extra-radical arbuscular mycorrhizal hyphae: a review. Plant and Soil 226, 199 –210. Koide, R.T. and Kabir, Z. (2000) Extraradical hyphae of the mycorrhizal fungus Glomus intraradices can hydrolyse organic phosphate. New Phytologist 148, 511–517. Kroehler, C. J., Antibus, R.K. and Linkins. A.E. (1988) The effects of organic and inorganic phosphorus concentration on the acid phosphatase activity of ectomycorrhizal fungi. Canadian Journal of Botany 66, 750 –756. Lassen, S.F., Breinholt, J., Østergaard, P.R., Brugger, R., Bischoff, A., Wyss, M. and Fuglsang, C.C. (2001) Expression, gene cloning, and characterization of five novel phytases from four basidiomycete fungi: Peniophora lycii, Agrocybe pediades, a Ceriporia sp., and Trametes pubescens. Applied and Environmental Microbiology 67, 4701–4707. Li, M., Osaki, M., Rao, I.M. and Tadano, T. (1997) Secretion of phytase from the roots of several plant species under phosphorus-deficient conditions. Plant and Soil 195, 161–169. Liu, Q., Loganathan, P., Hedley, M. J. and Skinner, M.F. (2004) The mobilisation and fate of soil and rock phosphate in the rhizosphere of ectomycorrhizal Pinus radiata seedlings in an allophanic soil. Plant and Soil 264, 219–229. Liu, Q., Loganathan, P. and Hedley, M. J. (2005) Influence of ectomycorrhizal hyphae on phosphate fractions and dissolution of phosphate rock in rhizosphere soils of Pinus radiata. Journal of Plant Nutrition 28, 1525–1540. Lung, S.-C. and Lim, B.L. (2006) Assimilation of phytate-phosphorus by the extracellular phytase activity of tobacco (Nicotiana tabacum) is affected by the availability of soluble phytate. Plant and Soil 279, 187–199. Lung, S.-C., Chan, W.-L., Yip, W., Wang, L., Yeung, E.C. and Lim, B.L. (2005) Secretion of beta-propeller phytase from tobacco and Arabidopsis roots enhances phosphorus utilisation. Plant Science 169, 341–349. Lynch, J.P. (2005) Root architecture and nutrient acquisition. In: Bassirirad, H. (ed.) Nutrient Acquisition by Plants: An Ecological Perspective. Springer, Berlin, Germany, pp. 147–183. Macfall, J., Slack, S.A. and Iyer, J. (1991) Effects of Hebeloma arenosa and phosphorus fertility on root acid phosphatase activity of red pine (Pinus resinosa) seedlings. Canadian Journal of Botany 69, 380–383. Martin, J.K. (1970) Organic phosphate compounds in water extracts of soils. Soil Science 109, 362–375. Martin, J.K. (1973) The influence of rhizosphere microflora on the availability of 32P-myo-inositol hexaphosphate phosphorus to wheat. Soil Biology and Biochemistry 5, 473–483. Martin, J.K. and Cartwright, B. (1971) The comparative plant availability of 32P myo-inositol hexaphosphate and KH232PO4 added to soils. Communications in Soil Science and Plant Analysis 2, 375–381. 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, 503–514. 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. Midgley, D. J., Chambers, S.M. and Cairney, J.W.G. (2004) Inorganic and organic substrates as sources of nitrogen and phosphorus for multiple genotypes of two ericoid mycorrhizal fungal taxa from Woollsia pungens and Leucopogon parviflorus (Ericaceae). Australian Journal of Botany 52, 63–71. Miller, S.S., Liu, J.Q., Allan, D.L., Menzhuber, C. J., Fedorova, M. and Vance, C.P. (2001) Molecular control of acid phosphatase secretion into the rhizosphere of proteoid roots from phosphorus-stressed white lupin. Plant Physiology 127, 594–606.

258


Mousain, D., Bousquet, N. and Polard, C. (1988) Comparison of phosphatase activities in ectomycorrhizal homobasidiomycetes cultured in vitro. European Journal of Forest Pathology 18, 299–309. Mudge, S.R., Smith, F.W. and Richardson, A.E. (2003) Root-specific and phosphate-regulated expression of phytase under the control of a phosphate transporter promoter enables Arabidopsis to grow on phytate as a sole phosphorus source. Plant Science 165, 871–878. Oberson, A. and Joner, E. J. (2005) Microbial turnover of phosphorus in soil. In: Turner, B.L., Frossard, E. and Baldwin, D.S. (eds) Organic Phosphorus in the Environment. CAB International, Wallingford, UK, pp. 133–164. Olsson, P.A., Jakobsen, I. and Wallander, H. (2002) Foraging strategies and resource allocation of mycorrhizal fungi in a patchy environment. In: van der Heijden, M. and Sanders, I. (eds) Mycorrhizal Ecology. Springer, Heidelberg, Germany, pp. 93–116. 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. Perez-Moreno, J. and Read, D. J. (2000) Mobilization and transfer of nutrients from litter to tree seedlings via the vegetative mycelium of ectomycorrhizal plants. New Phytologist 145, 301–309. Pierzynski, G.M., McDowell, R.W. and Sims, J.T. (2005) Chemistry, cycling and potential movement of inorganic phosphorus in soils. In: Sims, J.T. and Sharpley, A.N (eds) Phosphorus, Agriculture and the Environment. American Society of Agronomy, Madison, Wisconsin, pp. 53–86. Raghothama, K.G. (1999) Phosphate acquisition. Annual Reviews of Plant Physiology and Plant Molecular Biology 50, 665–693. Raghothama, K.G. (2005) Phosphorus and plant nutrition: an overview. In: Sims, J.T. and Sharpley, A.N. (eds) Phosphorus: Agriculture and the Environment. American Society of Agronomy, Madison, Wisconsin, pp. 355–378. Rausch, C. and Bucher, M. (2002) Molecular mechanisms of phosphate transport in plants. Planta 216, 23–37. Ravnskov, S., Larsen, J., Olsson, P.A. and Jakobsen, I. (1999) Effects of various organic compounds growth and phosphorus uptake of an arbuscular mycorrhizal fungus. New Phytologist 141, 517–524. Richardson, A.E. (2001) Prospects for using soil microorganisms to improve the acquisition of phosphorus by plants. Australian Journal of Plant Physiology 28, 897–906. 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. and Hayes, J.E. (2000) Phosphomonoesterase and phytase activities of wheat (Triticum aestivum L.) roots and utilisation of organic phosphorus substrates by seedlings grown in sterile culture. Plant, Cell and Environment 23, 397–405. Richardson, A.E., Hadobas, P.A. and Hayes, J.E. (2001a) Extracellular secretion of Aspergillus phytase from Arabidopsis roots enables plants to obtain phosphorus from phytate. The Plant Journal 25, 641–649. Richardson, A.E., Hadobas, P.A., Hayes, J.E., O’Hara, C.P. and Simpson, R.J. (2001b) Utilization of phosphorus by pasture plants supplied with myo-inositol hexaphosphate is enhanced by the presence of soil microorganisms. Plant and Soil 229, 47–56. Richardson, A.E., George, T.S., Hens, M. and Simpson, R.J. (2005) Utilisation of soil organic phosphorus by higher plants. In: Turner, B.L., Frossard, E. and Baldwin, D.S. (eds) Organic Phosphorus in the Environment. CAB International, Wallingford, UK, pp. 165–184. Rogers, H.T., Pearson, R.W. and Pierre, W.H. (1940) Absorption of organic phosphorus by corn and tomato plants and the mineralizing action of exo-enzyme systems of growing roots. Soil Science Society of America Proceedings 5, 285–291. Ryan, P.R., Delhaize, E. and Jones, D.L. (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.


259

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.

260


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


261

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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 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)


Sample origin and type

De Groot and Golterman (1993)

a

Proportion of the total organic phosphorus.

267

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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.)


269

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.)


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

271

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


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.

References Andersen, J.M. (1975) Influence of pH on release of phosphorus from lake sediments. Archiv für Hydrobiologie 76, 1725–1732. Bedford, M.R. (2000) Exogenous enzymes in monogastric nutrition: their current value and future benefits. Animal Feed Science and Technology 86, 1–13. Benson, R.L., McKelvie, I.D., Hart, B.T., Truong, Y.B. and Hamilton, I.C. (1996) Determination of total phosphorus in waters and wastewaters by on-line UV/thermal induced digestion and flow injection analysis. Analytica Chimica Acta 326, 29–39. 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. Boon, P. (1989) Organic matter degradation and nutrient regeneration in Australian freshwaters. 1. Methods for exoenzyme assays in turbid aquatic environments. Archiv für Hydrobiologie 115, 339–359. Boström, B., Jansson, M. and Forsberg, C. (1982) Phosphorus release from lake sediments. Archiv für Hydrobiologie Beihefte Ergebnisse der Limnologie 18, 5–59.


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Brearley, C.A. and Hanke, D.E. (1996a) Inositol phosphates in the duckweed Spirodela polyrhiza L. Biochemical Journal 314, 215–225. Brearley, C.A. and Hanke, D.E. (1996b) Metabolic evidence for the order of addition of individual phosphate esters in the myo-inositol moiety of inositol hexakisphosphate in the duckweed Spirodela polyrhiza L. Biochemical Journal 314, 227–233. Cade-Menun, B.J. and Preston, C.M. (1996) A comparison of soil extraction procedure for 31P NMR spectroscopy. Soil Science 161, 770–785. Caraco, N.F., Cole, J. J. and Likens, G.E. (1989) Evidence for sulphate-controlled phosphorus release from sediments of aquatic systems. Nature 341, 316–318. Celi, L., Lamacchia, S., Marsan, F.A. 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., Ajmore-Marsan, F. and Barberis, E. (2001) Effects of pH and electrolytes on inositol hexaphosphate interaction with goethite. Soil Science Society of America Journal 65, 753–760. Clarkin, C.M., Minear, R.A., Kim, S. and Elwood, J.W. (1992) An HPLC postcolumn reaction system for phosphorus-specific detection in the complete separation of inositol phosphate congeners in aqueous samples. Environmental Science and Technology 26, 199–204. Cooper, J.E., Early, J. and Holding, A. J. (1991) Mineralization of dissolved organic phosphorus from a shallow eutrophic lake. Hydrobiologia 209, 89–94. Cosgrove, D. J. (1980) Inositol Phosphates: Their Chemistry, Biochemistry and Physiology. Elsevier, Amsterdam, The Netherlands. 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. Einsele, W. (1936) Ueber die Beziehungen des Eisenkreislaufs zum Phosphatekreislauf im eutrophen See. Archiv für Hydrobiologie 29, 664–686. Eisenreich, S. J. and Armstrong, D.E. (1977) Chromatographic investigations of inositol phosphate esters in lake waters. Environmental Science and Technology 11, 497–501. Espinosa, M., Turner, B.L. and Haygarth, P.M. (1999) Preconcentration and separation of trace phosphorus compounds in soil leachate. Journal of Environmental Quality 28, 1497–1504. Gardolinski, P.C.F.C., Worsfold, P. J. and McKelvie, I.D. (2004) Seawater induced release and transformation of organic and inorganic phosphorus from river sediments. Water Research 38, 688–692. Golovan, S.P., Meidinger, R.G., Ajakaiye, A., Cottrill, M., Wiederkehr, M.Z., Barney, D. J., Plante, C., Pollard, J.W., Fan, M.Z., Hayes, M.A., Laursen, J., Hjorth, J.P., Hacker, R.R., Phillips, J.P. and Forsberg, C.W. (2001) Pigs expressing salivary phytase produce low-phosphorus manure. Nature Biotechnology 19, 741–745. Golterman, H.L. (1998) The distribution of phosphate over iron-bound and calcium-bound phosphate in stratified sediments. Hydrobiologia 364, 75–81. Golterman, H.L. (2001) Phosphate release from anoxic sediments or ‘What did Mortimer really write?’ Hydrobiologia 450, 99–106. Golterman, H., Paing, J., Serrano, L. and Gomez, E. (1998) Presence of and phosphate release from polyphosphates or phytate phosphate in lake sediments. Hydrobiologia 364, 99–104. Graneli, E. and Sundback, K. (1985) The response of planktonic and microbenthic algal assemblages to nutrient enrichment in shallow coastal waters, southwest Sweden. Journal of Experimental Marine Biology and Ecology 85, 253–268. Herbes, S.E., Allen, H.E. and Mancy, K.H. (1975) Enzymatic characterization of soluble organic phosphorus in lake water. Science 187, 432–434. Holdren, G.C. and Armstrong, D.E. (1980) Factors affecting phosphorus release from intact lake sediment cores. Environmental Science and Technology 14, 79–87. Hong, J.K. and Yamane, I. (1980) Examination of the conventional method and a proposal for a new method of determining inositol phosphate in the humic acid fraction. Soil Science and Plant Nutrition 26, 497–505. Inhulsen, D. and Niemeyer, R. (1978) Inositol phosphates from Lemna minor L. Zeitschrift für Pflanzenphysiologie 88, 103–116. 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.

276

I.D. McKelvie

Juanpere, J., Perez-Vendrell, A.M. and Brufau, J. (2004) Effect of microbial phytase on broilers fed barley-based diets in the presence or not of endogenous phytase. Animal Feed Science and Technology 115, 265–279. Kérouel, R. and Aminot, A. (1996) Model compounds for the determination of organic and total phosphorus dissolved in natural waters. Analytica Chimica Acta 318, 385–390. L’Annunziata, M.F. (1975) The origin and transformations of the soil inositol phosphate isomers. Soil Science Society of America Proceedings 39, 377–379. Leytem, A.B., Turner, B.L. and Thacker, P.A. (2004) Phosphorus composition of manure from swine fed low-phytate grains: evidence for hydrolysis in the animal. Journal of Environmental Quality 33, 2380–2383. Lijklema, L. (1977) The role of iron in the exchange of phosphate between water and sediments. In: Golterman, H.L. (ed.) Interactions between Sediments and Fresh Water. Dr W. Junk, BV, The Hague, The Netherlands, pp. 313–317. Littlewood, I.G. and Marsh, T. J. (2005) Annual freshwater river mass loads from Great Britain, 1975–1994: estimation algorithm, database and monitoring network issues. Journal of Hydrology 304, 221–237. Livingstone, D., Khoja, T.M. and Whitton, B.A. (1982) Laboratory studies on phosphorus sources for an Upper Teesdale Calothrix. British Phycological Journal 17, 235. Livingstone, D., Khoja, T.M. and Whitton, B.A. (1983) Influence of phosphorus on physiology of a hair-forming blue-green alga (Calothrix parietina) from an upland stream. Phycologia 22, 345–350. 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. McKelvie, I.D. (2005) Separation, preconcentration and speciation of organic phosphorus in environmental samples. In: Turner, B.L., Frossard, E. and Baldwin, D.S. (eds) Organic Phosphorus in the Environment. CAB International, Wallingford, UK, pp. 1–20. McKelvie, I.D., Hart, B.T., Cardwell, T. J. and Cattrall, R.W. (1993) Speciation of dissolved phosphorus in environmental samples by gel filtration and flow-injection analysis. Talanta 40, 1981–1993. McKelvie, I.D., Hart, B.T., Cardwell, T. J. and Cattrall, R.W. (1995) Use of immobilized phytase and flow injection for the determination of dissolved phosphorus species in natural waters. Analytica Chimica Acta 316, 277–289. McKercher, R.B. and Anderson, G. (1968) Content of inositol penta- and hexaphosphates in some Canadian soils. Journal of Soil Science 19, 47–55. Mortimer, C.H. (1941) The exchange of dissolved substances between mud and water in lakes. Journal of Ecology 29, 280–329. Mortimer, C.H. (1942) The exchange of dissolved substances between mud and water in lakes. Journal of Ecology 30, 147–201. Nanny, M.A. and Minear, R.A. (1994) Organic phosphorus in the hydrosphere. Characterization via 31P Fourier transform nuclear magnetic resonance. In: Baker, L.A. (ed.) Environmental Chemistry of Lakes and Reservoirs. American Chemical Society, Washington, DC, pp. 161–191. Oku, O. and Kamatani, A. (1995) Phospholipid in plankton samples from Tokyo Bay and Sagami Bay. Nippon Suisan Gakkaishi 61, 588–595. 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. Potman, W. and Lijklema, L. (1983) Inositol-phosphates: hydrolysis, stability and analysis. Water Research 17, 411–414. Rippey, B. (1977) The behaviour of phosphorus and silicon in undisturbed core of Lough Neagh sediments. In: Golterman, H.L. (ed.) Interactions between Sediments and Fresh Water. Dr W. Junk, B.V., The Hague, The Netherlands, pp. 349–353. Roberts, R.M. and Loewus, F.A. (1968) Inositol metabolism in plants. VI. Conversion of myoinositol to phytic acid in Wolffiella floridana. Plant Physiology 43, 1710–1716. Roden, E.E. and Edmonds, J.W. (1997) Phosphate mobilization in iron-rich anaerobic sediments: microbial Fe(III) oxide reduction versus iron-sulfide formation. Archiv für Hydrobiologie 139, 347–378. Rozan, T.F., Traillfert, M., Trouwborst, R.E., Glazer, B.T., Ma, S., Herszage, J., Valdez, L.M., Price, K.S. and Luther, G.W. (2002) Iron–sulfur–phosphorus cycling in the sediments of a shallow coastal bay: implications for sediment release and benthic microalgal blooms. Limnology and Oceanography 47, 1346–1354. Sharpley, A.N. and Withers, P. J.A. (1994) The environmentally sound management of agricultural phosphorus. Fertilizer Research 39, 133–146.


277

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.


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

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