myo-inositol metabolism in plants

Plant Science 150 (2000) 1–19

Review

myo-Inositol metabolism in plants

Frank A. Loewus a,*, Pushpalatha P.N. Murthy b

a Institute of Biological Chemistry, Washington State Uni6ersity, Pullman, WA 99164-6340, USAb Department of Chemistry, Michigan Technological Uni6ersity, Houghton, MI 49931, USA

Received 3 June 1999; received in revised form 19 July 1999; accepted 19 July 1999

Abstract

The multifunctional position supplied by myo-inositol is emerging as a central feature in plant biochemistry and physiology. Inthis critique, attention is drawn to metabolic aspects and current assessment is made of manifold ways in which myo-inositol andits metabolic products impact growth and development. The fact that a unique enzyme, common to all eukaryotic organismswhere such assessment has been undertaken, controls conversion of D-glucose-6-P to 1L-myo-inositol-1-P provides a useful pointof departure for this brief metabolic survey. Some aspects such as biosynthesis, phosphate and polyphosphate ester hydrolysis, andO-methylation of myo-inositol have captured the consideration of molecular biologists, yet other aspects including oxidation,conjugation, and transfer to phospholipids remain virtually untouched from this viewpoint. Here, an attempt is made to enlist newinterest in all facets of myo-inositol metabolism and its place in plant biology. © 2000 Elsevier Science Ireland Ltd. All rightsreserved.

Keywords: Galactinol; Galactopinitol; Glycosylphosphatidylinositol; Glycosylinositolphosphorylceramide; IAA-MI conjugates; myo-Inositol;1L-myo-Inositol-1-P; Ins(3)P1 synthase; MI kinase; MI monophosphatase; MI oxidation pathway; O-Methyl Inositols; Ononitol; Phosphatidyli-nositols; Phytic acid; Pinitol; Raffinose

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1. Introduction

Isolation of myo-inositol (MI) from muscle ex-tract by Scherer in 1850 led to 80 years of intenseinterest on the natural occurrence, properties,derivatives, and stereoisomers of the cyclitols.Then, discovery that MI functioned as a growthfactor for certain microorganisms and as a re-quirement for growth of certain mutant forms ofyeast prompted fresh interest in its biochemicaland biological features. Soon it became apparentthat MI played a central role in growth and devel-

opment [1]. This was especially true in plant biol-ogy where molecular entities containing orutilizing MI were involved in structure and func-tion [2]. Fig. 1 summarizes this information bycategorizing specific products of MI metabolismwith particular interest to plant biologists and byidentifying avenues of inquiry which may lead to abetter appreciation of this unique molecule and itsposition in plant science. Colored backgrounds inFig. 1 attempt to provide a sense of related func-tions while avoiding the confusion of ‘metabolicmapping’.

Conversion of D-glucose-6-P to 1L-MI-1-P con-stitutes the first committed step in MI biosynthesis[1–3]. Metabolic processing of MI beyond biosyn-thesis produces other stereo-forms of inositol andleads to a host of functional roles, all of whichrequire this unique cyclitol. These include:

Abbre6iations: ABA, abscisic acid; IAA, indole-3-acetic acid; MI,myo-inositol; Ins(3)P1, 1L-myo-inositol-1-P; InsP6 or MI-P6, phyticacid; MIOP, myo-inositol oxidation pathway; SNOP, sugar nucle-otide oxidation pathway.

* Corresponding author. Tel.: +1-509-335-3413; fax: +1-509-335-7643.

E-mail address: [email protected] (F.A. Loewus)

0168-9452/00/$ - see front matter © 2000 Elsevier Science Ireland Ltd. All rights reserved.

PII: S 0 1 6 8 -9452 (99 )00150 -8

F.A. Loewus, P.P.N. Murthy / Plant Science 150 (2000) 1–192

� Cycling of 1L-MI-1-P and free MI by MI phos-phatase and MI kinase [4,5].

� Oxidation of free MI to D-glucuronic acid withits subsequent role in biogenesis of uronosyland pentosyl units of pectin, hemicelluloses,and related structures in plant cell walls [1,2,6].

� Esterification of MI to form auxin (IAA) estersand their glycosides [7,8].

� Conjugation of free MI with UDP-D-galactoseto form galactinol, the galactosyl donor for

biosynthesis in the raffinose and galactopinitolseries of oligosaccharides [9,10].

� Isomerization and methylation of MI and otherisomeric (scyllo-, chiro-, muco-, and neo-) inosi-tols to form O-methyl inositols (sequoyitol,bornesitol, quebrachitol, pinitol, ononitol, etc.)which participate in stress-related responses,storage of seed products, and production ofinositol-glycosides such as pinitol-galactosides[9–13].

Fig. 1. Functional roles of myo-inositol in plant metabolism.

F.A. Loewus, P.P.N. Murthy / Plant Science 150 (2000) 1–19 3

Fig. 2. myo-Inositol.

If one takes biosynthesis as a basis of assign-ment and traces the origin from D-glucose 6-phos-phate, then clockwise assignment preservesbiosynthetic relatedness. Unhappily, recent ad-vances involving key roles for MI polyphosphatesinvolved in signal transduction (where addition orloss of a single phosphate may alter assignmentfrom D- to L- or vice-versa) have created confusionfor those unfamiliar with rules of cyclitol nomen-clature. The upshot is a tentative agreement by theInternational Union of Biochemistry to relax rulesof nomenclature so that 1L-MI-1-P, the product ofmyo-inositol-1-phosphate synthase, may be desig-nated 1L-MI(1)P1, 1D-MI(3)P1, or simply, Ins(3)P1

where the symbol Ins signifies MI with counter-clockwise numbering from 1D. Thus, 1D-MI(1,4,5)P3, an important physiological signalgenerated during phosphatidylinositol-4,5-bisphos-phate metabolism, becomes simply Ins(1,4,5)P3.More detailed discussion of the stereochemistry ofMI and its phosphate esters is found in [14] andon the Internet at http://www.chem.qmw.ac.uk/iubmb/nomenclature/.

3. MI biosynthesis

3.1. E6idence for cyclization of D-glucose to MI

Although the D-gluco configuration inherent inMI was recognized by Maquenne as early as 1887and the proposition that D-glucose-6-phosphatecyclized to form Ins(3)P1 enzymatically was ad-vanced by H.O.L. Fischer in 1945, unequivocalevidence for conservation of the 6-carbon chain ofD-glucose during cyclization to MI did not appearuntil 1962 [20,21].

The experimental approach involved recovery oflabeled MI from a D-[1-14C]glucose-labeled parsleyleaf followed by administration of this labeled MIto detached immature strawberry fruits where itwas utilized as a carbon source for pectin biosyn-thesis [20,22]. Carbon-14 was recovered in D-galac-turonosyl and L-arabinosyl residues of pectinwhich upon radioanalysis revealed 79% of thelabel in carbon 1. Distribution of 14C in sucrose-derived D-glucose, pectin-derived D-galacturonateand L-ascorbic acid from the parsley leaf also had\80% of their 14C in carbon 1. In other words,about 80% of the 14C in these products of D-(1-14C)glucose metabolism remained at the original

Fig. 3. Conventions for numbering substituents in myo-inosi-tol.

� Biosynthesis of phytic acid (MI-P6) and phyticacid pyrophosphates [14–16].

� Metabolic recycling of products of phytic acidhydrolysis during phytase-mediated phytic aciddephosphorylation [14–16].

� Biosynthesis of phosphatidylinositol, its poly-phosphates, and precursors of MI polyphos-phate-specific signal transduction [1,2,14,17].

� Glycosylated-phosphatidylinositol and glycosy-lated-inositolphosphorylceramide [18,19].

2. Nomenclature

The nomenclature of inositols has been an on-going source of confusion and conflict for decades.MI is a meso compound with a plane of symmetrythat rotates the structure about C2 and C5 as fixedpositions (Fig. 2). The remaining four carbonatoms consist of two prochiral pairs, C1=C3 andC4=C6. If the carbon ring is numbered clock-wise, as shown by numbers inside the ring, assign-ment of a single substituent on carbon 1 is 1L.Conversely, if the carbon ring is numbered coun-terclockwise as shown by numbers external to thering, assignment is 1D. 1L-MI-1-P, the product of1L-myo-inositol-1-phosphate synthase (E.C.5.5.1.4), is a good example of the dilemma this choicecreates (Fig. 3).


site of labeling. Some redistribution of 14C be-tween terminal carbons is normal during triose/hexose phosphate metabolism.

This analytical approach provided profound in-sight into three aspects of MI metabolism, namelyMI biosynthesis, the MI oxidation pathway, andphytic acid biosynthesis. It provided evidence forcyclization of the carbon chain of D-glucose toform MI. It revealed an alternative biosyntheticpathway to uronic acid and pentose constituentsof plant cell wall polysaccharides quite indepen-dent of the one involving UDP-D-glucose dehy-drogenase. Finally, it supplied stereochemicalevidence for the putative initial phosphorylatedintermediate leading to phytic acid, a major formof phosphate storage in plants.

3.2. Mechanism of MI biosynthesis

Biosynthetic conversion of D-glucose to free MIinvolves three enzymatic steps (Fig. 4). Step B,cyclization of D-glucose-6-P to Ins(3)P1, is the firstcommitted step in MI biosynthesis. Step C, loss ofphosphate, releases free MI. Overall, this schemeconstitutes the sole pathway of MI biosynthesis incyanobacteria, algae, fungi, plants, and animalsand occupies a central role in their cellularmetabolism.

Cyclization of D-glucose-6-phosphate toIns(3)P1 is irreversible. This process also highlightsa dilemma in nomenclature since D-glucose isnumbered clockwise about the pyranose ring whileconventional numbering of MI inverts the num-bering of its inherent D-gluco configuration.

Ins(3)P1 synthase appears to be a highly con-served enzyme [23]. Functionally, this conversionof D-glucose 6-P to Ins(3)P1 involves three sub-steps (Fig. 5):� NAD+-coupled oxidation of carbon 5 of D-

glucose-6-P.� Aldol condensation between carbon 1 and car-

bon 6 of 5-keto-D-glucose-6-P (D-xylo-5-hexulose-6-P).

� NADH-catalyzed reduction of 2-myo-inosose-1-P (D-2,4,6/3,5-pentahydroxy-cyclohexane-2-Pto yield Ins(3)P1.

Specific points of interest regarding Ins(3)P1 syn-thase include:� Preference for the b-anomeric form of glucose-

6-P.� Enzyme-bound NAD (removable by charcoal

treatment to generate an inactive apo-enzyme).� NAD+-catalyzed oxidation at carbon 5 of glu-

cose-6-P (substep 1) to yield an enzyme-bound5-keto-glucose-6-P with hydride ion transferfrom glucose-6-P to the pro-S position of car-bon 4 on the nicotinamide moiety of NAD+

(Kinetic studies indicate a sequential reactionwith NAD+ adding first. There is a distinctisotope effect in removal of hydrogen fromcarbon 5 of glucose-6-P.).

� Base-catalyzed cyclization (substep 2).—Oxy-gen at carbon 5 is retained. The pro-R hydro-gen is removed from carbon 6 while the pro-Shydrogen is retained. The second intermediatewithin brackets is 2-myo-inosose-1-P (D-2,4,6/3,5-pentahydroxycyclohexanone-2-P). For addi-tional details, see [24].

Fig. 4. Conversion of D-glucose to MI: (A) Hexokinase, EC 2.7.1.1; (B) Ins(3)P1 synthase, EC 5.5.1.4; (C) MI monophosphatase,EC 3.1.3.25.

Fig. 5. Enzymatic mechanism of 1L-MI-1-phosphate synthase (a.k.a. Ins(3)P1 synthase).


� Stereospecific oxidation of enzyme-boundNADH (substep 3) including transfer of itspro-S hydride ion to the si face of the carbonylgroup to generate Ins(3)P1.

� An undisturbed phosphate–carbon bond.Hydrolysis of Ins(3)P1 by a specific MI

monophosphatase [4] completes overall conversionof D-glucose to free MI.

3.3. Biochemical and physiological aspects ofIns(3)P1 synthase

The structural gene INO1 for Ins(3)P1 synthasewas first isolated from the yeast Saccharomycescere6isiae by Donahue and Henry [25]. Subsequentstudies involving ino1 mutations in MI auxotrophsprovided insight into regulatory control processes[3,26]. Transcripts with homology to this genehave been obtained from several plant sources andare summarized in these cited references [3,23].The first such plant gene for Ins(3)P1 synthase tobe characterized was tur1, a cDNA from the duck-weed, Spirodela polyrrhiza, which was rapidly andspatially up-regulated during an ABA-inducedmorphogenic response [27]. This effect was local-ized to stolon tissue that connects the developingturion to the node of the mother frond. Theauthors considered several possible scenarios inwhich MI synthesis might play a role. These in-cluded phytic acid accumulation, lipid synthesis,an ABA signal transducing mechanism involvingphosophoinositide metabolites, altered flux of MImetabolites into the cell wall and/or an auxin-linked cell elongation involving auxin conjugates,and induced response to stress involving methylethers of inositol. Of these possibilities, an effecton the nature of cell wall structures appeared mostinteresting. Since S. polyrrhiza was not yet trans-formable at the time of these studies, Smart andFlores [28] generated transgenic Arabidopis plantsover-expressing Ins(3)P1 synthase TUR1 cDNAfrom S. polyrrhiza and found these plants to con-tain elevated Ins(3)P1 synthase activity with con-comitant four-fold increase in endogenous MI.Comparison of transgenic to wild-type plants re-vealed no significant differences in whole plantgrowth habit, expansion growth, germination rate,flowering time, stem thickness, in vitro rootgrowth and germination/survival on high salt orlow temperature regimes. A four-fold increase inendogenous MI may have been insufficient to

trigger gross differences in growth or developmentalthough compositional differences have been ob-served in lily pollen germinated in media that wassupplemented with MI ranging from 0.3 to 2.8mM [29].

In algae and plants, both cytosolic and chloro-plastic forms of Ins(3)P1 synthase have been iso-lated and characterized [30]. Although thebiochemical and kinetic parameters of these twoforms do not differ significantly between eachother or from other cytosolic Ins(3)P1 synthasespreviously described [3], the native cytosolic formis homotrimeric while the native chloroplasticform is homotetrameric. Interestingly, a cyanobac-terium, Spirulina platensis, included in the citedstudy, contained only one cytosolic homote-trameric form as anticipated by the endosymbionttheory for a cyanobacterial origin of plastids[3,31,32].

Analysis of an INO1-like transcript (termedINPS1 by the authors [23]) from salt-stressedMesembryanthemum crystallinum (ice plant) re-vealed a diurnal fluctuating increase in mRNAduring the light period that could be coordinatedwith the gene encoding MI-O-methyltransferase,an enzyme methylating MI to D-ononitol which isepimerized to D-pinitol. D-Pinitol accumulates insalt-stressed M. crystallinum plants and is consid-ered to be the principal osmoregulator. Compara-ble experiments with Arabidopsis thalianatranscripts failed to produce this effect and theauthors conclude that there is probably no stress-mediated induction of Inps 1 mRNA in Arabidop-sis, an observation similar to that made by Smartand Flores [28]. Salt-tolerant varieties of ricegrown in a NaCl environment exhibited a photore-sponsive enhancement of chloroplast and cytosolicIns(3)P1 synthase activity [33]. The authors specu-late on the possible role of free MI as an osmolytein the chloroplast through coordinate activationand/or induced expression of Ins(3)P1 synthaseand MI monophosphatase.

Keller et al. [34] obtained a full-length cDNAfrom potato epidermal tissue that encodedIns(3)P1 synthase (termed StIPS-1). RNA blotanalysis revealed the highest StIPS-1 transcriptlevels in photosynthetic tissues but much lowerlevels in roots and tubers. Light greatly elevatedStIPS-1 transcript levels but drought stress had noeffect. When antisense StIPS-1 transformantswere tested, it was found that their leaves had


strongly reduced levels of MI, galactinol, andraffinose. These plants also showed distinct mor-phological aberrations including decreased overalltuber yield. These findings highlight broad bio-chemical and physiological effects brought on byaltering Ins(3)P1 synthase activity in potato andpossibly most other plant species.

Quite recently, Yoshida and co-workers [35]isolated a cDNA clone, pRINO1, from rice (Oryzasati6a L.) callus suspension cultures that is highlyhomologous to Ins(3)P1 synthase from yeast andplants. Its transcript appears in the apical regionof globular-stage embryos 2 days after anthesisand strong signals were detected in the scutellumand aleurone layer after 4 days. Phytate-contain-ing particles or globoids appeared in the sametissues at 4 days, coinciding with the RINO1transcript. This study demonstrates that Ins(3)P1

synthase is probably the first committed step inphytic acid biosynthesis although a complemen-tary process involving salvage of MI by MI kinaseremains untested.

In studies just reviewed, genes encoding Ins(3)P1

synthase are variously identified as INO1 (Saccha-romyces cere6isiae), TUR1 (Spirodela polyrrhiza),StIPS-1 (Solanum tubersum), INPS1 (Mesem-bryanthemum crystallinum), pRINO1 (Oryza sa-ti6a), etc. Deduced amino acid sequences obtainedfrom these plant-derived sources are quite similar.In the interest of consistency, a common term likeINPS1 as proposed by Ishitani et al. [23] seemsdesirable.

3.4. Concerning free MI

Dephosphorylation of Ins(3)P1 constitutes thesole de novo route to free MI in plants. All othersources derive from salvage mechanisms involvingrecovery of free MI from other metabolic MI-con-taining products. Free MI is generally regarded asa ubiquitous constituent of plant tissues and insome species, notably Actinidia arguta, (kiwifruit),MI is the major ‘sugar’ constituent (60–65%) dur-ing the first 20–30 days after anthesis [36]. Stress-related aspects of MI accumulation have beennoted repeatedly [1,2,6,23,37,38] but specific bio-chemical and molecular details are needed. In fact,accumulation of free MI may be a more universalphenomenon in life forms than generally realized.In overwintering ladybird beetles (Creatomegillaundecimnotata), free MI, functioning as a possible

cryoprotectant, increases more than four-fold(from 2.5 to 11 mg/mg wet weight) during wintermonths [39].

A relatively specific alkaline, magnesium-depen-dent phosphatase (MI monophosphatase, EC3.1.3.25) from lily pollen hydrolyzes Ins(3)P1, itsenantiomer Ins(1)P1, and at a somewhat lowerrate, Ins(2)P1, to free MI [40,41]. Animal tissuescontain a similar MI monophosphatase [42] butIns(2)P1 which is substituted in the axial positionis not a substrate although it does act as a compet-itive inhibitor in the case of bovine brain enzyme.There is need to revisit this matter of substratespecificity since most plant studies are dated in thisregard.

Gillaspy et al. [4] cloned three MI monophos-phatase activities from tomato (LeIMP). All iso-forms were lithium ion-sensitive over aconcentration range similar to that exhibited byhuman MI monophosphatase. When labeled anti-sense RNA probes were used to follow mRNAaccumulation of these isoforms at different devel-opmental stages and in different organs, LeIMP1mRNA accumulation was greatest in light-grownseedlings, flowers, young and mature green fruit(decreasing as fruit proceeded to the breakerstage), and callus tissue. LeIMP3 mRNA wasdetected in the same tissues and had a muchhigher response in the shoot apex. LeIMP2mRNA levels were significantly lower than theother two isoforms. Consideration of apparentdifferences in the expression patterns of these threeisoforms, prompted the suggestion that their activ-ities functioned within different cell types or spa-tially distinct cellular compartments. Evidence tosupport spatially regulated expression has sincebeen presented [43]. Given the diverse demands forfree MI as outlined in Fig. 1 and the ancestralhistory of plant organelles, such a suggestion isquite possible. Clearly, there is need for moreinformation on localization of MI monophos-phatase within the plant cell.

Inhibition of MI monophosphatase by lithiumions is especially noteworthy. At 0.1 mM Li+,bacterial protein extracts, each expressing one ofthe LeIMP gene products, had less than 20% ofmaximal activity for removing phosphate from[14C]MI-1-P [4]. Although phytotoxicity of Li+ hasbeen studied for decades, efforts to pinpoint itsspecific targets in plant cells have yielded onlymeager results [44]. Beyond its inhibitory effect on


Fig. 6. Cell wall polysaccharide biogenesis via the myo-inositol oxidation pathway (bold font) and the sugar nucleotide oxidationpathway.

MI monophosphatase, little is known regardingits impact on availability of free MI for numer-ous biosynthetic and regulatory requirements(Fig. 1). Lithium ion delayed initiation of DNAsynthesis and cell division when introduced intosynchronized Catharanthus roseus cell cultures, acondition largely prevented when MI was in-cluded in the medium. Preparations of MImonophosphatase from C. roseus were inhibited80% by 10 mM Li+ [45]. Use of Li+ inhibitionas a tool for studying modulation of free cellularMI appears to be a viable option, one whichmay also alter other metabolic pathways con-nected to a demand for free MI such as its roleas substrate for MI kinase (EC 2.7.1.64). Curi-ously, little attention has been given to this latterenzyme which is present in plants, animals andmicroorganisms [46]. Its product, Ins(3)P1, hasthe same configurational structure as that pro-duced by Ins(3)P1 synthase [5]. While one mightregard this recycling of MI back into a pool ofIns(3)P1 as a salvage mechanism, it fails to takeinto consideration any localization of these en-zymic activities or temporal demands during de-velopment. Together, Ins(3)P1 synthase and MIkinase constitute ways in which Ins(3)P1 isformed from D-glucose-6-P or free MI in plants.The former enzyme is biosynthetic while the lat-ter must rely on sources that generate free MIfrom MI monophosphatase or other MI-conju-gated forms. Unresolved are temporal and spa-tial patterns of synthase and kinase duringgrowth and development.

4. myo-Inositol oxidation pathway

Substantial experimental support for a MIoxidation pathway (MIOP) in plants hasaccumulated during the past 35 years since thispathway was first proposed [22] with over 50 papersfrom Loewus’s laboratory alone addressing thistopic. The presence of a MIOP has beendemonstrated in a wide variety of plant tissuesincluding strawberry fruit, parsley leaf, lily floralparts and pollen, pear pollen, cultured sycamoreand rice callus, corn root-tip, duckweed,germinating and developing wheat, pine pollen,rubber latex serum, and algae.

The MIOP (Fig. 6) involves cyclization ofD-glucose-6-P to Ins(3)P1, loss of phosphate to formMI, oxidation of MI to D-glucuronic acid,phosphorylation at carbon 1 (a configuration), andconversion by uridylyl transferase to UDP-D-glucuronic acid. Alternatively, D-Glucose-6-P isconverted to UDP-D-glucose, which undergoesoxidation to UDP-D-glucuronic acid, a processtermed the sugar nucleotide oxidation pathway(SNOP) [1,2,6,21]. In Fig. 6, bold borders contraststeps of the MIOP and its metabolic products fromthose of the SNOP.

Both UDP-D-glucuronic acid and its product ofdecarboxylation, UDP-D-xylose, strongly inhibitNAD+-dependent UDP-D-glucose dehydrogenase[47]. The requirement for NAD+ as well as kineticrestraints imposed by product inhibition areimportant considerations when invoking the SNOPfor UDP-D-glucuronic acid metabolism. Neither ofthese effects appears in the MIOP. This hassignificant implications in that inhibition of UDP-


D-glucose oxidation will leave the MIOP as theprincipal pathway to UDP-D-glucuronate and itsproducts. Relative fluxes in demands on UDP-D-glucose coupled to the inhibitory effect of UDP-D-xylose on oxidation of UDP-D-glucose may wellallow MIOP to play a major role in hexose/pen-tose metabolism, cell wall pectin and hemicelluloseformation, and starch synthesis [48–50]. When thehydrogen isotope effect, which occurs duringIns(3)P1 synthase-catalyzed conversion of D-[5-3H]glucose-6-P to [2-3H]MI, was utilized to com-pare the functional role of the sugar nucleotideand MI oxidation pathways in germinating lilypollen [51], 3H/14C ratios of glucosyl and galactur-onosyl residues from amyloglucosidase/pectinasehydrolysates strongly supported the view that con-version of glucose into galacturonic acid residuesof pectin used the MIOP. Recently, UDP-D-glu-cose dehydrogenase has been isolated and purifiedfrom soybean nodules [52] and cloned from soy-bean cell suspension cultures [53]. The expressionpattern of the latter was studied at selected devel-opmental stages. Results suggest that this enzymehas a key regulatory role in production of hemicel-lulosic precursors. Access to this cloned gene forUDP-D-glucose dehydrogenase together withthose for Ins(3)P1 synthase and MI monophos-phatase should provide the tools needed in futurestudies to dissect expression patterns of the twopathways at critical stages of growth anddevelopment.

5. Indole-3-acetic acid (IAA) conjugates of MIand its glycosides

Within the family of IAA-conjugates found inplants are a number of IAA-esters that includeIAA-MI and IAA-MI with galactosyl or ara-binosyl substituents [7,8]. Biosynthesis of IAA-MIis a two step reaction mechanism involving UDP-glucose and free MI as substrates:IAA+UDP-glucoseB.

\1-O-IAA-glucose+UDP1-O-IAA-glucose+myo-inositolB.

\IAA-myo-inositol+glucose

Addition of sugar residues as 5-b-O- sub-stituents follows esterification. These esters aregenerally regarded as inactive ‘storage’ formswhereby plants cope with excess auxin production.

Alternately, such structures might facilitate trans-port of auxin within the plant. Apart from the factthat free MI is required for IAA-MI production,little is known regarding the metabolic fate of thismoiety. During tropic response of a plant to anasymmetric stimulus, there is asymmetric distribu-tion of IAA. Assuming that the source of thishormone is an IAA conjugate such as IAA-MI,then the fate of MI may well be related to itscapacity to undergo oxidation via the MIOP andenter into cell wall constituents of pectin and/orhemicellulose. Such studies have yet to beundertaken.

6. Inositols and their methyl ethers

6.1. Galactosyl inositols (raffinose andgalactopinitol series)

There are nine isomeric structures for inositol(Fig. 7). Eight isomers are diastereomeric, one ofwhich is enantiomeric. Naturally occurring inosi-tols and/or their methyl ethers in plants includemyo-, scyllo-, muco-, neo-, +chiro-, and −chiro-inositol [54–56]. Only myo-inositol (MI) is synthe-sized de novo from hexose phosphate by MI1-phosphate synthase and MI monophosphatase.Production of other isomeric inositols involvesmetabolic processing of MI. For many years fol-lowing their isolation and characterization, inter-est in these inositols was limited largely to theirchemistry [57] and to their occurrence, which insome cases appeared to be species-specific [58]. Animportant physiological role for MI emerged withthe discovery that galactinol (O-a-D-galactopyra-nosyl (1�3)-Ins) functioned as galactosyl donorfor biosynthesis of the trisaccharide, raffinose andfor higher galactosyl homologues that are involvedin phloem transport, seed development, seed desic-cation, and numerous stress-related responses ofplants [9–12,59]. Subsequent studies uncovered O-methyl inositol-based galactosides containingononitol (4-O-methyl Ins) and pinitol (1D-4-O-methyl-chiro-inositol) that are presumably in-volved in galactosyl storage and transfer [6,7,60].A biosynthetic relationship between galactinol andgalactosylononitol emerged when it was discov-ered that stachyose synthase (EC 2.4.1.67),purified to homogeneity from mature seed ofadzuki beans (Vigna angularis), contained the fol-lowing activities [13]:


� Stachyose synthase (galactinol donor),raffinose + galactinol� stachyose + myo - ino-sitol

� An exchange reaction, myo-[3H]inositol+galactinol� [3H]galactinol+myo-inositol

� Stachyose synthase (galactosylononitol donor),raffinose + galactosylononitol � stachyose +ononitol

� Galactosylononitol synthase–ononitol+galac-tinol�galactosylononitol+myo-inositolRe-analysis of the structure of a galactosylonon-

itol previously isolated from adzuki bean [61] re-veals it to be identical to the same galactosyl-ononitol involved in raffinose and stachyosebiosynthesis [62].

Galactinol synthase utilizes UDP-D-galactose asits galactosyl donor. In enzyme preparations ofrice bean (Vigna umbellata), galactinol is a re-quired galactosyl donor for galactosylononitol andthis, in turn, donates galactosyl moities toraffinose and stachyose. Only galactinol functionsas galactosyl donor to galactosylononitol [12]. Astudy of ononitol biosynthesis and accumulationin V. umbellata during drought stress found thatmethylation of MI appears to correlate with stem-localized methyltransferase activity and a signifi-cant rise in ononitol in the leaves [63]. Themethyltransferase is specific for the 4-hydroxyl ofIns (a.k.a., 6-hydroxyl of 1L-MI) and shows noactivity with other naturally occurring isomericinositols and O-methyl inositols [63].

6.2. Stress-related responses in6ol6ing inositol orO-methyl inositol

A broad variety of naturally occurring plantconstituents contribute to osmotic regulation inplants that have been exposed to stressful environ-ments [64]. Included among these are the inositolsand their O-methyl ethers. The term ‘stress’ is usedhere to refer to such abiotic factors as drought,heat, cold, salinity, pollutants, and reactive oxygenspecies. Brief overviews are found in several stud-ies involving inositols and stress tolerance [9–11,23,33,35,37,38,56,65,66].

The halophytic ice plant (Mesembryanthemumcrystallinum) is a model for studies involving in-duced crassulacean metabolism. Plants irrigatedwith 400 mM sodium chloride accumulate pinitolwhich eventually constitutes over two-thirds of thesoluble carbohydrate fraction and approximately10% of the dry weight [67]. The osmotic adjust-ment of this plant is generally attributed to itshigh level of pinitol. Bohnert and his colleagues[11] have studied the biosynthetic pathway thatleads to pinitol accumulation with the hope thatthis effort will lead to a better understanding ofdrought and salt tolerance. The pathway involvesthree enzymic steps beyond hexose phosphate(Fig. 8).

They found two Inps-like transcripts encodingIns(3)P1 synthase, the first enzyme in the pathway,with deduced amino acid sequences nearly identi-

Fig. 7. Isomeric inositols.


Fig. 8. Biochemical conversion of myo-inositol to pinitol. Bracket indicates theoretical intermediate.

cal to Ins(3)P1 synthase from other plants. Undersalinity stress, the Inps1RNA up-regulated five-foldor more and free MI accumulated ten-fold [23].The second step, an O-methyl transferase, methy-lates C4 of MI. Transgenic tobacco plants bearingIMT1, the O-methyl transferase from ice plant,were found to accumulate ononitol and providebetter protection under drought and salt-stressconditions than wild-type plants [68]. The enzymecatalyzing step three, epimerization of C1 of onon-itol, has yet to be examined but it is assumed thatthis step is not rate-limiting. Immunocytology andsolute measurements [69] led to discovery of in-creased phloem transport of MI accompanied byincreased transport of Na+ and inositol to leavesof ice plant under stress. It was found thatseedlings of ice plants, which are not salt-tolerant,developed patterns of gene expression and polyolaccumulation observed in mature salt-tolerantplants and that MI-enhanced Na+ uptake andtransport increased [38]. From these data it isproposed that a Na+/MI symport might exist thatpromotes Na+ uptake in the ice plant. This is anovel idea which, as the authors point out, may bea general mechanism of controlling Na+ uptake inglycophytes. It also offers new clues regarding theosmoregulatory role of O-methyl inositols.

It is safe to assume that research on inositol-linked, stress-related processes in plants is still inits pioneering stages. There is an interesting obser-vation that sap collected during the dormant pe-riod (winter and early spring) from sugar maple(Acer saccharum) is rich in quebrachitol (1L-2-O-methyl-chiro-inositol), ranging from 4–6% of total

solids. Yet when trees break dormancy virtuallyno quebrachitol remains in the sap [70]. Currentstudies on the role of cyclitols as stable organicosmolytes in trees may be one explanation [65].

7. Structure and conformation of phytic acid

In 1872, Pfeffer showed that subcellular parti-cles in wheat endosperm contained a calcium/mag-nesium salt of organic phosphate. Two structureswere proposed for phytic acid and it took over 50years to resolve the structural issue. In 1908, Neu-berg proposed a structure that contained threecyclic pyrophosphate moieties and in 1914, Ander-son proposed a structure in which the six hydroxylgroups on MI are esterified with orthophosphatemoieties [15,57,71–73]. NMR spectroscopy finallyresolved the choice in favor of the latter in 1969[74].

In foods and feed, phytic acid reduces thebioavailibility of inositol, phosphorus, and essen-tial minerals by forming non-assimilable salt com-plexes which lead to detrimental nutritional effectson human and animals [75,76]. Moreover, disposi-tion of this unabsorbed phytate creates environ-mental phosphorus contamination, an agriculturalsituation that has prompted governmental legisla-tion in Europe and the USA [76]. To meet thischallenge, agricultural interests have actively en-couraged development of plant cultivars with re-duced phytic acid as well as related methods ofmodifying foods and feeds to render their phytatecontent more nutritional [77].


Cellular roles proposed for phytic acid includinginhibition of protein phosphatases and subsequentmodulation of calcium channel activity, attenua-tion of endocytosis, and inhibition of clathrinassembly [78,79]. Here, further work is needed tofully confirm these functions.

Although the orientation of the phosphategroups in phytic acid was established, there wasmuch debate about the conformation adopted[71,80]. 1ax/5eq and 5ax/1eq (Fig. 9) differ signifi-cantly in overall molecular shape and the orienta-tion of polar groups. Consequently, the twoconformations exhibit significant differences inchelating ability, including interaction withproteins. Therefore, the conformational preferenceof phytic acid in different environments is criticalto understanding the chemistry and biochemistryof phytic acid. Assignment of conformation(s) forphytic acid [71] has only recently been fully re-solved. 1H-NMR spectroscopic methods [80] en-abled Barrientos and Murthy [81] to show that MIadopts the sterically favorable 1ax/5eq conforma-tion (one phosphate axial and five equatorial) inpH range 1.0–9.0 and the sterically hindered 5ax/1eq conformation above pH 9.5 (Fig. 9). BetweenpH 9.0–9.5 (the pKa region of the three leastacidic protons) both conformations are in dynamicequilibrium. Dynamic NMR indicates that theactivation energy for the conformational inversionprocess is �54.890.8 kJ/mole (compared to thatfor ring inversion of cyclohexane, about 45 kJ/mole) [82]. Inversion to the 5ax/1eq form is facili-tated by complexation with metal ions whichreduces electrostatic repulsion and thereby stabi-lizes the sterically hindered, dodeca-anionic form.Stabilization of the 5ax/1eq form is influenced bythe size of the counter ion. Alkali metals Na+,K+, Rb+ and Cs+ (with hydrated radii less than

or equal to 2.76 A, ) stabilize the 5ax/1eq formwhereas Li+ ion (hydrated radii 3.4 A, ) does notstabilize that form. In the presence of larger coun-ter ions such as tetramethylammonium and tetra-butylammonium ions, the presence of the stericallyhindered 5ax/1eq form is not observed, thus indi-cating that complexation with counter ions is es-sential for ring inversion of phytic acid [82].

NMR spectroscopy also indicated that the con-formational preferences of individual isomers atdifferent pH’s are dictated by structural featuresunique to the isomer such as the number of phos-phate moieties and the regiochemical and stereo-chemical arrangement of the phosphates [81].Ins(1,2,3,4,6)P5 adopts the 1ax/5eq form in the pHrange 1.0–9.0 and above 9.5 both 1ax/5eq and5ax/1eq forms exist in dynamic equilibrium; theexclusive presence of the 5ax/1eq form is notobserved. Inositol phosphates containing less than5 phosphates showed no proclivity to undergo ringinversion to the sterically hindered form [81].

Molecular modeling studies were carried outusing ab initio, semi-empirical and force fieldmethods (MacroModel V6.0 and Gaussian 94), forboth the 1ax/5eq and 5ax/1eq conformations ofphytic acid in the fully protonated and dodeca-an-ionic state [82]. Molecular modeling calculationswere consistent with NMR results in aqueous so-lution. Interestingly, calculations predicted thatthe relative stability of the two conformations isthe same in vacuum and aqueous solution,namely, in the fully protonated state the stericallyfavourable 1ax/5eq form of phytic acid is morestable than the 5ax/1eq form and that in thedodeca-anionic state the sterically hindered 5ax/1eq form is more stable than the 1ax/5eq form[82].

8. Biosynthesis of Phytic Acid

Although the presence of phytic acid in plantcells has been known for over a century, attemptsto determine its biosynthesis in whole plants, plantorgans, subcellular organelles, and cell cultures[14,77,78,83], have only recently seen signs of pro-gress. Evidence suggests that phytic acid biosyn-thesis occurs in cisternal endoplasmic reticulumand the product is subsequently deposited inphytin granules [84]. Investigations in duckweed,Spirodela polyrhiza [16] and the slime mold, Dic-

Fig. 9. Conformational structures of phytic acid: (1) the1ax/5eq form (pH 1–9). (2) The 5ax/1eq form (\ pH 9.5).


tyostelium [85,86], provided strong evidence forsequential phosphorylation of Ins(3)P1.

Spirodela polyrrhiza :Ins(3)P1�Ins(3,4)P2�Ins(3,4,6)P3

�Ins(3,4,5,6)P4�Ins(1,3,4,5,6)P5

�InsP6

Dictyostelium :Ins(3)P1�Ins(3,6)P2�Ins(3,4,6)P3

�Ins(1,3,4,6)P4�Ins(1,3,4,5,6)P5

�InsP6

Both pathways begin with Ins(3)P1 which makesbiosynthetic sense in that Ins(3)P1 is product ofboth Ins(3)P1 synthase and MI kinase. The closerelationship between Ins(3)P1 formation andphytic acid biosynthesis in developing seeds isrevealed in the elegant work of Yoshida et al. [35].As discussed earlier in regard to Ins(3)P1 biosyn-thesis, in situ hybridization of developing ricegrains seeds showed that the transcript forIns(3)P1 synthase (RINO1) first appeared in theupper half of the embryo two days after anthesis,increased for the next five days and then graduallydecreased. Phytin-containing globoids first ap-peared in the scutellum and the aleurone layerfour days after anthesis and increased gradually inboth tissues. The appearance of globoids coincidedwith the localization of RINO1 transcripts therebysuggesting that enhanced Ins(3)P1 formationdrives phytic acid biosynthesis. It is interestingthat phosphorylation at the 1 position of Ins(P)n

occurs quite late in both biosynthetic pathwayssuggesting that the second messenger Ins(1,4,5)P3

pathway and the phytic acid biosynthetic pathwaydo not intersect in these tissues. In contrast, phos-pholipase C-triggered production of Ins(1,4,5)P3

and subsequent conversion to phytic acid, which isinvolved in mRNA transport, have been shown inyeast [87]. In all pathways proposed, the 2 positionof Ins(P)n is the last position to bephosphorylated.

9. Hydrolysis of phytic acid:

The enzymes responsible for phytic acid hydrol-ysis are phytases, a special class of phosphatasesthat catalyze the sequential hydrolysis of phyticacid [14,15]. The sequence of hydrolysis by acidphytases was first established by Tomlinson and

Ballou [88,89]. Their methods for determining thestructures of intermediate inositol phosphates arestill widely used. Based on the position of the firstphosphate hydrolyzed, two classes of acid phytasesare recognized, the 6-phytase (EC 3.1.3.26) andthe 3-phytase (EC 3.1.3.8) [15]. Both hydrolyzephytic acid to Ins(2)P1. The X-ray crystal structureof a 3-phytase from Aspergillus niger has beendetermined [90]. The structure consists of a largea/b-domain, which shows structural similarity to ahigh molecular weight acid phosphatase in rats,and a smaller a-domain. The active site containsthe amino acid sequence, RHGXRXP, common inmany acid phosphatases, and a cluster of basicamino acid residues that could facilitate binding ofthe negatively charged phytic acid at the activesite. The enzyme did not contain bound substrate;therefore a model for substrate binding and cata-lytic activity was proposed by the authors. In themodel, the conserved histidine (His 59) was in-volved in nucleophilic attack at the 3-phosphate.

An unusual constitutive alkaline phytase that ispresent in lily pollen and seeds [91,92] initiatesaction by first removing the 5-phosphate of phyticacid. Subsequent hydrolytic steps remove phos-phate from the 4 and 6 position to yield Ins(1,2,3)P3 as the final product. This final product,Ins(1,2,3)P3, has been shown to inhibit iron-cata-lyzed free radical formation by chelating iron [93–95]. The presence of multiple phytases withdiffering specificity, pH optima, and biochemicalproperties in wheat bran and lily pollen suggeststhat hydrolysis of phytic acid is under the controlof multiple phytases. We are probably entering anew phase of research which will involve under-standing the physiological importance of the mul-tiple phytases and the biological roles of inositolphosphates produced by them.

10. Pyrophosphorylated inositol phosphates

Recently, inositol phosphates containing one ormore pyrophosphate groups, have been detected inslime mold and mammalian cells [78,79]. Com-pounds with a pyrophosphate group at the 1 or 5position of phytic acid (pyrophosphoinositol pen-takisphosphate, PP-InsP5) or two pyrophosphategroups tentatively placed at 1 and 5 or at 5 and 6positions (bispyrophosphoinositol tetrakisphos-phate, [PP]2-InsP4) have been identified (Fig. 10).A phytic acid-kinase, which can convert the phos-


Fig. 10. Pyrophosphorylated inositol polyphosphates. (A) 5-PP-InsP5 and (B) 1,5-[PP]2-InsP4.

phate group at the 5 position to a high energypyrophosphate group, and a PP-InsP5 kinase,which can convert a second phosphate to a py-rophosphate group, have been isolated [96]. Thesehigh-energy pyrophosphates are not metabolicallystatic. They exhibit rapid turnover through theaction of phosphatases and kinases. Estimates arethat 30–50% of the cellular phytic acid cyclesthrough these pyrophosphorylated derivatives ev-ery hour [97]. The potential involvement of py-rophosphorylated inositol phosphates in signaltransduction and calcium metabolism is suggestedby the observation that in a muscle cell line, theconcentration of [PP]2-InsP4 declined rapidly(70%) by activation of the cAMP-dependent path-way and that sigargin, which increases cytoplasmiccalcium by depleting calcium stores in endoplas-mic reticulum, reduced the concentration of [PP]-InsP5 and [PP]2-InsP4 by 50% (reviewed in Shears[79]). Such tantalizing bits of information indicatethat these metabolically-active, high energy phos-phates could play multiple roles in phosphate andcalcium metabolism and suggest that further workis necessary for a better understanding of theircellular roles.

11. Phosphatidylinositol and its polyphosphates

Due to the critical role that phosphoinositidesplay in signal transduction, this area of inositolchemistry has been actively investigated and itscurrent status in both animals and plants has beencomprehensively reviewed [98]. Phosphatidylinosi-tol (PtdIns) and its phosphorylated derivatives,PtdIns-4-phosphate (PtdInsP) and PtdIns-4,5-bis-phosphate (PtdInsP2), are present in plant tissues[98] but their functions as extracellular signalsremains an equivocal issue [99]. (As used in thisreview, the abbreviation PtdIns refers to PtdInswith a MI head group. If the head group is otherthan MI, the isomeric inositol will be specified.) In

plants, the biosynthesis of PtdIns via the CDP-dia-cylglycerol pathway is well established [98] (Fig.11), as is subsequent phosphorylation of PtdIns byspecific kinases to PtdInsP and PtdInsP2. Morerecently, the presence of the 3-substitutedpolyphosphoinositides, PtdIns(3)P1 and Pt-dIns(3,4)P2, in plant cells has been documented[98].

Besides variability in the position and numberof phosphate groups on the inositol ring, anothersource of structural heterogeneity in phosphoinosi-tides has been the presence of isomeric inositols.Although MI is the predominant form in themajority of inositol-containing compounds, thepresence of scyllo-inositol-containing PtdIns (des-ignated PtdscylloI) has been found in plant cellsand chiro-inositol-containing PtdIns (designatedPtdchiroI) in animal cells. Isomeric phosphoinosi-tides could be synthesized by unique CDP-DG:inositol 3-phosphatidyltransferase, also calledPtdIns synthase, such as CDP-DG:scyllo-inositol3-phosphatidyl transferase (Fig. 11, step 4) or byexchange of the head group, MI for scyllo-inositolby PtdIns:scylloI phosphatidyltransferase, alsocalled the exchange enzyme (Fig. 11, step 5). Atpresent there is evidence for both pathways. Invitro studies in barley aleurone cells have indicatedthe presence of CDP-DG:scyllo-inositol trans-ferase activity. Taken together, in vivo and in vitrostudies suggest that PtdscylloI is biosynthesized bythe CDP-DG:scyllo-inositol transferase pathway[100]. Investigations in mammalian cells [101] andsoybean seedlings [102] suggested that PtdchiroI issynthesized via PtdIns either by the operation of ahead group exchange enzyme or an epimerase. Inthis context, it is important to note the cloningstudies in mammalian cells which indicated that asingle polypeptide exhibited both the synthase andCMP-dependent exchange activities [103]. There-fore, further work is required to demonstrateclearly that head group transfer activity is notinvolved in the biosynthesis of PtdscylloI in barleyaleurone cells.


12. Glycosyl-phosphatidylinositol andGlycosyl-inositolphosphorylceramide

The discovery that glycosylated lipid moleculesanchor proteins to cell membranes and that boththe lipid anchors and the proteins are involved incellular response to environmental stimuli hasopened up an exciting area of research [104–106].In plant cells, reports of the involvement of twosuch anchors, a glycosyl–PtdIns (GPI)-anchorednitrate reductase in blue light-stimulated nitrateuptake [107,108] and stimulation of glycosyl–ce-ramide-anchored alkaline phosphatase in low-phosphate medium [18,109], suggest that they mayplay important roles in plants as well.

The structure of the lipid moiety that anchorsenzymes to the hydrophobic membranes has beeninvestigated in a number of cells and structuralinformation gleaned from these studies revealsthat the GPI anchors have conserved and variablestructural moieties [18,104–106] (Fig. 12). In the

conserved core structure, the 6-hydroxyl of the Insmoiety is glycosylated with a tetrasaccharide chaincontaining one glucosamine and three mannoseunits. The third mannose is connected to a phos-phorylated ethanolamine, the amine group ofwhich forms an amide bond with the C-terminalend of the protein. Variability in the lipid portionincludes the presence of a 1-alkyl-2-acyl glycerolor a ceramide unit in place of the diacylglycerol(see below) and variation in the fatty acid compo-sition. Structural heterogeneity in the hydrophilicportion of the molecule includes esterification of ahydroxyl at carbon 2 on the MI moiety with along chain fatty acid, the presence of chiro-inosi-tol, galactose molecules glycosidically linked to thecore mannose groups, and two or threeethanolamine molecules. Biosynthesis of GPI an-chors involves the sequential glycosylation of Pt-dIns in a step-wise manner and post translationalmodification of the protein with the preformedGPI anchor [105].

Fig. 11. Scheme for biosynthesis of phosphatidylinositols with different head groups.


Fig. 12. Structures of (A) glycosyl-phosphatidylinositol and (B) glycosyl-inositolphosphorylceramide.

Employing methods developed for animal cells,Tischner and his colleagues showed that the inosi-tol-containing lipid anchor of nitrate reductaseincludes a diacylglycerophosphatidyl-MI moiety[107,108]. These methods include (i) release of theprotein from membrane vesicles by the action ofPtdIns-specific phospholipase C, (ii) decrease inthe hydrophobicity of the released protein, (iii) invivo labeling of the anchored protein by[3H]ethanolamine, and (iv) cross reactivity withmonoclonal antibody raised against the GPI-an-chor of Trypanosoma GPI-anchored protein[107,108]. However, the lipid anchor of alkalinephosphatase in Spirodela exhibits chemical reactiv-ity that is not consistent with the presence of aGPI unit [18,109]. These characteristics include,incorporation of myo-[3H]inositol, [3H]ethano-lamine, [3H]myristic and [3H]palmitic acid intolipid-bound enzyme, resistance to cleavage ofprotein by PtdIns-specific phospholipase C, andhydrolysis of fatty acid units under strong acidand alkaline conditions but not under mild alkali.On the basis of these chemical characteristics, theauthors have tentatively identified that the lipidmoiety is a ceramide [18,109]. Thus the parentlipid must be a glycosylated inositolphosphorylce-ramide, also called glycophosphosphingolipid [19](Fig. 12). The presence of glycophosphosphin-golipid, namely lipids containing the inositolphos-

phorylceramide group, in seeds from cotton,peanut, corn and soybeans has been known sincethe 1960s as a result of the pioneering work byCarter et al. [19]. However, our knowledge of thebiosynthesis, localization, and biological roles ofsphingolipids is limited. In light of the continuallyexpanding role of lipids in signal transduction andthe realization that sphingolipids mediate cellgrowth and differentiation in animal cells, therecent discoveries [18,104] of the involvement ofinositol-containing glycosyl-lipids in nutrient as-similating processes has opened up an excitingarea for future research [110].

13. Concluding Remarks

A beguiling simplicity shrouds a staggeringcomplexity of reactions, pathways and physiologi-cal consequences arising from the unique biochem-ical properties of MI. Inherently, MI bears theD-gluco configuration. Conformational responsesto physical and chemical changes associated withsubstituent groups present a bewildering array ofreactivities. From a stereochemical viewpoint, thissingle molecule poses prochiral choices at four ofits six hydroxylated sites. Superimposed on thesechoices is the nature of substituent group(s). Theseconsiderations must be included in any effort to


understand the central position of MI in plantmetabolism and its multifunctional role. To reachsuch goals requires more knowledge of the regula-tion and control of MI biosynthesis, the selectivenature of substituent groups, the interplay of iso-meric forms, the phosphorylative process, cellulardeposition of fully phosphorylated MI as phytate,recall of phytic acid during critical stages of seedgermination and seedling growth as well as infor-mation regarding metabolic redistribution of theenergy and chemical fragments released duringcatabolism of phytic acid. It also entails a muchgreater effort to explore the properties of phos-phatidylinositols and their polyphosphates as po-tential signals in plant cells. Here, qualitiescommon to plant and animal systems must besorted out from those expressly unique to plants.Last, but certainly not least, the relationship be-tween oxidation of MI and cell wall biogenesisneeded to be revisited. As Trethewey, Krotzky &Willmitzer [111] point out, ‘‘…the time is ripe formetabolic profiling and…this will emerge as acentral aspect of plant functional genomics in thenext decade.’’. The task of garnering this informa-tion lies ahead.

Acknowledgements

For many years, studies in FAL’s laboratoryhave been supported by grants from the NationalInstitutes of Health. The authors also wish toacknowledge Dr. Søren Rasmussen, Risø NationalLaboratory, Denmark, whose 1997 Workshop onPhytate and Phytases in Plants provided the im-petus for this review. The authors thank thosewho reviewed this manuscript for their usefulcomments.

References

[1] F.A. Loewus, M.W. Loewus, myo-Inositol: Its biosyn-thesis and metabolism, Annu. Rev. Plant Physiol. 34(1983) 137–161.

[2] D.J. Morre, W.F. Boss, F.A. Loewus (Eds.), InositolMetabolism in Plants, Wiley–Liss, 393 p, New York,1990.

[3] A.L. Majumder, M.D. Johnson, S.A. Henry, 1L-myo-Inositol-1-phosphate synthase, Biochim. Biophys. Acta1348 (1997) 245–256.

[4] G.E. Gillaspy, J.S. Keddie, K. Oda, W. Gruissem, Plantinositol monophosphatase is a lithium-sensitive enzyme

encoded by a multigene family, Plant Cell 7 (1995)2175–2185.

[5] M.W. Loewus, K. Sasaki, A.L. Leavitt, L. Munsell,W.R. Sherman, F.A. Loewus, Enantiomeric form ofmyo-inositol-1-phosphate produced by myo-inositol-1-phosphate synthase and myo-inositol kinase in higherplants, Plant Physiol. 70 (1982) 1661–1663.

[6] F. Loewus (Ed.), Biogenesis of Plant Cell Wall Polysac-charides, Academic Press, New York, 1973, p. 379.

[7] J.P. Slovin, R.S. Bandurski, J.D. Cohen, Control ofhormone synthesis and metabolism 1. Auxins, in: P.J.J.Hooykaas (Ed.) Biochemistry and Molecular Biology ofPlant Hormones, Elsevier, Amsterdam (in press).

[8] J.D. Cohen, J.P. Slovin, Recent research advances con-cerning indole-3-acetic acid metabolism, in: K. Palme,R. Walden, J. Schell (Eds.), Mechanism of Action ofPlant Hormones, Springer, Berlin (in press).

[9] M. Horbowicz, R.L. Obendorf, Seed desiccation toler-ance and storability: Dependence on flatulence-produc-ing oligosaccharides and cyclitols (review and survey,Seed Sci. Res. 4 (1994) 385–405.

[10] R.L. Obendorf, Oligosaccharides and galactosyl cycli-tols in seed desiccation (review update), Seed Sci. Res. 7(1997) 63–74.

[11] H. Bohnert, R.G. Jensen, Strategies for engineeringwater-stress tolerance in plants, Trends Biotechnol. 14(1996) 89–97.

[12] T. Peterbauer, M. Puschenreiter, A. Richter,Metabolism of galactosylononitol in seeds of Vignaumbellata, Plant Cell Physiol. 39 (1998) 334–341.

[13] T. Peterbauer, A. Richter, Galactosylononitol andstachyose synthesis in seeds of Adzuki bean, PlantPhysiol. 117 (1998) 165–172.

[14] P.P.N. Murthy, Insoitol phosphates and theirmetabolism in plants, in: B.B. Biswas, S. Biswas (Eds.),myo-Inositol Phosphates, Phosphoinositides, and SignalTransduction, Subcellular Biochemistry Series, vol. 26,Plenum Press, New York, 1996, pp. 227–255.

[15] D.J. Cosgrove, Inositol Phosphates: Their Chemistry,Biochemistry and Physiology, Elsevier, Amsterdam,1980, p. 191.

[16] C.A. Brearley, D.E. Hanke, Metabolic evidence for theorder of addition of individual phosphate esters to themyo–inositol moiety of inositol hexaphosphate in theduckweed Spirodela polyrhiza, Biochem. J. 314 (1996)227–233.

[17] G.G. Cote, R.C. Crain, Biochemistry of phosphoinosi-tides, Annu. Rev. Plant Physiol. Mol. Biol. 44 (1993)333–356.

[18] H. Nakazato, T. Okamoato, M. Nishikoori, K. Washio,N. Morita, K. Haraguchi, G.A. Thompson Jr., H.Okuyama, The glycosylphosphatidylinositol-anchoredphosphatase from Spirodela oligorrhiza is a purple acidphosphatase, Plant Physiol. 118 (1998) 1015–1020.

[19] R.A. Laine, T.C.-Y. Hsieh, Inositol-containing sphin-golipids, Methods Enzymol. 138 (1987) 186–195.

[20] F.A. Loewus, S. Kelly, Conversion of glucose to inosi-tol in parsley leaves, Biochem. Biophys. Res. Commun.7 (1962) 204–208.

[21] F.A. Loewus, Inositol metabolism and cell wall forma-tion in plants, Fed. Proc. FASEB 24 (1965) 855–863.


[22] F.A. Loewus, S. Kelly, E.F. Neufeld, Metabolism ofmyo-inositol in plants: Conversion to pectin, hemicellu-lose, D-xylose and sugar acids, Proc. Natl. Acad. Sci.USA 48 (1962) 421–425.

[23] M. Ishitani, A.L. Majumder, A. Bornhouser, C.B.Michalowski, R.C. Jensen, H.J. Bohnert, Coordinatetranscriptional induction of myo-inositol metabolismduring environmental stress, Plant J. 9 (1996) 537–548.

[24] M.E. Migaud, J.W. Frost, Elaboration of a generalstrategy for inhibition of myo-inositol 1-phosphate syn-thase: Active site interactions of analogues possessingoxidized reaction centers, J. Am. Chem. Soc. 118 (1996)495–501.

[25] T.F. Donahue, S.A. Henry, myo-Inositol-1-phosphatesynthase: Characteristics of the enzyme and identifica-tion of its structural gene in yeast, J. Biol. Chem. 256(1981) 7077–7085.

[26] M.J. White, J.M. Lopes, S.A. Henry, Inositolmetabolism in yeasts, Adv. Microbial Physiol. 32 (1991)1–51.

[27] C.C. Smart, A.J. Fleming, A plant gene with homologyto D-myo-inositol-3-phosphate synthase is rapidly andspatially up-regulated during an abscisic-acid-inducedmorphogenic response in Spirodela polyrrhiza, Plant J. 4(1993) 279–293.

[28] C.C. Smart, S. Flores, Overexpression of D-myo-inosi-tol-3-phosphate synthase leads to elevated levels ofinositol in Arabidopis, Plant Mol. Biol. 33 (1997) 811–820.

[29] M. Chen, F.A. Loewus, myo-Inositol metabolism inLilium longiflorum pollen, Plant Physiol. 59 (1977) 653–657.

[30] A. RayChaudhuri, N.C. Hait, S. DasGupta, T.J.Bhaduri, R. Deb, A.L. Majumber, l-myo-Inositol 1-phosphate synthase from plant sources: Characteristicsof the chloroplastic and cytosolic enzymes, Plant Phys-iol. 115 (1997) 727–736.

[31] D. Bhattacharya, L. Medlin, Algal phylogeny and theorigin of land plants, Plant Physiol. 116 (1998) 9–15.

[32] W.F. Doolittle, A paradigm gets shifty, Nature 392(1998) 15–16.

[33] A. RayChaudhuri, A.L. Majumber, Salinity-inducedenhancement of l-myo-inositol 1-phosphate synthase inrice (Oryza sati6a L.), Plant Cell Environ. 19 (1996)1437–1442.

[34] R. Keller, C.A. Brearley, R.N. Trethewey, B. Muller-Rober, Reduced inositol content and altered morphol-ogy in transgenic potato plants inhibited for1D-myo-inositol 3-phosphate synthase, Plant J. 16(1998) 403–410.

[35] K.T. Yoshida, T. Wada, H. Koyama, R. Mizobuchi–Fukuoka, S. Naito, Temporal and spatial patterns ofaccumulation of the transcript of myo-inositol-1-phos-phate synthase and phytin-containing particles duringseed development in rice, Plant Physiol. 119 (1999)65–72.

[36] K. Klages, H. Donnison, H. Boldingh, E. MacRae,myo-Inositol is the major sugar in Actinidia argutaduring enarly fruit development, Aust. J. Plant Physiol.25 (1998) 61–67.

[37] J. Bucker, R. Giderian, Accumulation of myo-inositolin Populus as a possible indication of membrane disinte-gration due to air pollution, J. Plant Physiol. 144 (1994)121–123.

[38] D.E. Nelson, M. Koukoumanos, H.J. Bohnert, myo-Inositol-dependent sodium uptake in ice plant, PlantPhysiol. 119 (1999) 165–172.

[39] V. Kostal, O. Nedved, P. S& imek, Accumulation of highconcentrations of myo-inositol in the overwintering la-dybird beetle Creatomegilla undecimnotata, Cryo Letters17 (1996) 267–272.

[40] M.W. Loewus, F.A. Loewus, myo-Inositol-1-phos-phatase from the pollen of Lilium longiflorum Thunb,Plant Physiol. 70 (1982) 765–770.

[41] S.C. Gumber, M/W. Loewus, F.A. Loewus,@ Furtherstudies on myo-inositol-1-phosphatase from the pollenof Lilium longiflorum Thunb., Plant Physiol. 76 (1984)40–44.

[42] L. Parthasarathy, R.E. Vadnal, R. Parthasarathy, C.S.Shyamala Devi, Biochemical and molecular propertiesof lithium-sensitive myo-inositol monophosphatase: Aminireview, Life Sci. 54 (1994) 1127–1142.

[43] J.A. Styer, W. Gruissem, G.A. Gillaspy, Spatially regu-lated expression of tomato inositol monophosphatasegenes, Plant Physiol. ASPP Meeting Suppl. (1999)Abst.149.

[44] C.E. Anderson, Lithium in plants, in: R.O. Bach, V.S.Gallicchio (Eds.), Lithium and Cell Physiology,Springer–Verlag, New York, 1990, pp. 24–46.

[45] T. Nishida, H. Kodama, A. Komamine, The effects oflithium chloride on the induction of proliferation ofcells in suspension cultures of Catharanthus roseus, J.Plant Physiol. 142 (1993) 184–190.

[46] P.D. English, M. Deitz, P. Albersheim, Myoinositolkinase: partial purification and identification ofproduct, Science 151 (1966) 198–199.

[47] D.S. Feingold, Aldo (and keto) hexoses and uronicacids, in: F.A. Loewus, W. Tanner, (Eds.) Plant Carbo-hydrates I, Encyclopedia of Plant Physiology, NewSeries, Vol. 13A, Springer Verlag, Berlin, 1982, pp.3–76.

[48] C-L. Rosenfield, C. Fann, F.A. Loewus, Metabolicstudies on intermediates in the myo-inositol oxidationpathway in Lilium longiflorum pollen: I. Conversion tohexoses, Plant Physiol. 61 (1978) 89–95.

[49] C.L. Rosenfield, F.A. Loewus, Metabolic studies onintermediates in the in the myo-inositol oxidation path-way in Lilium longiflorum pollen: II. Evidence for theparticipation of UDP-xylose and free xylose as interme-diates, Plant Physiol. 61 (1978) 96–100.

[50] C.L. Rosenfield, F.A. Loewus, Metabolic studies onintermediates in the in the myo-inositol oxidation path-way in Lilium longiflorum pollen: III. Polysaccharidicorigin of labeled glucose, Plant Physiol. 61 (1978) 101–103.

[51] M.W. Loewus, F.A. Loewus, The C-5 hydrogen isotopeeffect in myo-inositol 1-phosphate synthase as evidencefor the myo-inositol oxidation pathway, Carbohydr.Res. 82 (1980) 333–342.

[52] D.C. Stewart, L. Copeland, Uridine 5’-diphosphate-glu-cose dehydrogenase from soybean nodules, Plant Phys-iol. 116 (1998) 349–355.


[53] R. Tenhaken, O. Thulke, Cloning of an enzyme thatsynthesizes a key nucleotide-sugar precursor of hemicel-lulose biosynthesis from soybean: UDP-glucose dehy-drogenase, Plant Physiol. 112 (1996) 1127–1134.

[54] R. Mukherjee, E.M. Axt, Cyclitols from Croton celtidi-folius, Phytochemistry 23 (1984) 2682–2684.

[55] F.A. Loewus, Structure and occurrence of inositols inplants: see Ref. 2, pp. 1–11.

[56] M. Popp, N. Smirnoff, Polyol accumulation andmetabolism during water deficit, in: N. Smirnoff (Ed.),Environment and Plant Metabolism, BIOS Sci. Publ,Oxford, UK, 1995, pp. 199–215.

[57] T. Posternak, The Cyclitols, Holden-Day, San Fran-cisco, 1965, p. 431.

[58] V. Plouvier, Distribution of aliphatic polyols and cycli-tols, in: T. Swain (Ed.), Chemical Taxonomy, AcademicPress, New York, 1963, pp. 313–336.

[59] O. Kandler, H. Hopf, Oligosaccharides based on su-crose (sucrosyl oligosaccharides), in: F.A. Loewus, W.Tanner (Eds.), Encyclopedia of Plant Physiology, NewSeries, vol. 13A, Springer Verlag, 1982, pp. 349–383.

[60] T.M. Kuo, C.A. Lowell, T.C. Nelson, Occurrence ofpinitol in developing soybean seed tissues, Phytochem-istry 45 (1997) 29–35.

[61] T. Yasui, Identification of a new galactosyl cyclitolfrom seeds of Vigna angularis Ohwi et Ohashi (adzukibean), Agric Biol. Chem. 44 (1980) 2253–2255.

[62] A. Richter, T. Peterbauer, I. Brereton, Structure ofgalactosylononitol, J. Nat. Prod. 60 (1997) 749–751.

[63] W. Wanek, A. Richter, Biosynthesis and accumulationof D-ononitol in Vigna umbellata in response to droughtstress, Planta 197 (1995) 424–427.

[64] B.N. Timmerman, C. Steelink, F.A. Loewus (Eds.),Phytochemical Adaptations to Stress, Recent Advancesin Phytochemistry, vol. 18, Plenum Press, New York,1983.

[65] M. Popp, W. Leid, U. Bierbaum, M. Gross, T. Grosse-Schulte, S. Hams, J. Oldenettel, S. Schuler, J. Wiese,Cyclitols, stable osmotica in trees, in: H. Rennenberg,W. Eschrich, H. Ziegler (Eds.), Contributions to Mod-ern Tree Physiology, SPB Academic Publ, The Hague,1997, p. 270.

[66] N. Smirnoff, Q.J. Cumbes, Hydroxyl radical scavengingactivity of compatible solutes, Phytochemistry 28 (1989)1057–1060.

[67] M.J. Paul, W. Cockburn, Pinitol, a compatible solute inMesembryanthemum crystallinum L.?, J. Exper. Bot. 40(1989) 1093–1098.

[68] E. Sheveleva, W. Chmara, H.J. Bohnert, R.G. Jensen,Increased salt and drought tolerance by D-ononitolproduction in transgenic Nicotiana tabacum L, PlantPhysiol. 115 (1997) 1211–1219.

[69] D.E. Nelson, G Rammesmayer, H.J. Bohnert, Regula-tion of cell-specific inositol metabolism and transport inplant salinity tolerance, Plant Cell 10 (1998) 753–764.

[70] E.E. Stinson, C.J. Dooley, J.M. Purcell, J.S. Ard, Quer-bachitol (a new component of maple sap and syrup,Agric. Food Chem. 15 (1967) 394–397.

[71] R. Lasztity, L. Lasztity, Phytic acid in cereal technol-ogy, Adv. Cereal Sci. Technol. 10 (1990) 309–371.

[72] B.F. Harland, E.R. Morris, Phytate: A good or badfood component?, Nutrition Res. 15 (1995) 733–754.

[73] S.E. Rickard, L.U. Thompson, Interactions and biolog-ical effects of phytic acid, in: Antinutrients and Phyto-chemicals in Food, American Chemical SocietySymposium Series No. 662, Washington D.C., 1997, pp.294–312.

[74] L.F. Johnson, M.E. Tate, Structure of ‘phytic acids’,Can. J. Chem. 47 (1969) 63–73.

[75] D.S. Ertl, K.A. Young, V. Raboy, Plant genetic ap-proaches to phosphorous management in agriculturalproduction, J. Environ. Qual. 27 (1998) 299–304.

[76] D.T. Van der Molen, A. Breeuwsman, P.C.M. Boers,Agricultural nutrient losses to surface water in theNetherlands: Impact, strategies, and perspectives, J. En-viron. Qual. 27 (1998) 4–11.

[77] V. Raboy, P. Gerbasi, Genetics of myo-inositol phos-phate synthesis and accumulation, in: B.B. Biswas, S.Biswas (Eds.), Inositol Phosphates, Phosphoinositides,and Signal Transduction, Subcellular Biochemistry se-ries, vol. 26, Plenum Press, New York, 1996, pp. 257–285.

[78] S.B. Shears, Inositol pentskis- and hexakisphosphatemetabolism adds versatility to the actions of inositolpolyphosphates novel effects on ion channels andprotein traffic, in: B. Biswas, S. Biswas (Eds.), InositolPhosphates, Phosphoinositides, and Signal Transduc-tion, Subcellular Biochemistry series, vol. 26, PlenumPress, New York, 1996, pp. 187–226.

[79] S.B. Shears, The versatility of inositol phosphates ascellular signals, Biochim. Biophys. Acta 1436 (1998)49–67.

[80] L.R. Isbrandt, R.P. Oertel, Conformational states ofmyo-inositol hexakis (phosphate) in aqueous solution.A 13C NMR, 31P NMR, and Raman spectroscopicinvestigation, J. Amer. Chem. Soc. 102 (1980) 3144–3148.

[81] L.G. Barrientos, P.P.N. Murthy, Conformational stud-ies of myo-inositol phosphates, Carbohydr. Res. 296(1996) 39–54.

[82] A.T. Bauman, G.M. Chateauneuf, R.E. Brown, P.P.N.Murthy, Conformational inversion process in phyticacid: NMR spectroscopic and molecular modeling stud-ies, Tetrahedron Lett. 40 (1999) 4489–4492.

[83] J.J. Scott, F.A. Loewus, Phytate metabolism in plants,in: E. Graf (Ed.), Phytic acid: Chemistry and Applica-tions, Pilatus Press, Minneapolis, 1986, pp. 23–42.

[84] J.N.A. Lott, J.S. Greenwood, G.D. Batten, Mechanismsand regulation of mineral nutrient storage during seeddevelopment, in: J. Kigel, G. Galili (Eds.), Seed Devel-opment and Germination, Marcel Dekker, New York,1995, pp. 215–235.

[85] L.R. Stephens, R.F. Irvine, Stepwise phosphorylationof myo-inositol leading to myo-inositol hexakisphos-phate in Dictyostelium, Nature (London) 346 (1990)580–583.

[86] P.J.M. van Haastert, P. van Dijken, Biochemistry andgenetics of inositol phosphate metabolism in Dictyos-telium, FEBS Lett. 410 (1997) 39–43.

[87] J.D.York Odom, A.R. Murphy, R. Ives, E.B. Wente, Aphospholipase C-dependent inositol polyphosphate ki-


nase pathway required for efficient messenger RNAexport, Science 285 (1999) 96–100.

[88] R.V. Tomlinson, C.E. Ballou, myo-Inositol polyphos-phate intermediates in the dephosphorylation of phyticacid by phytases, Biochemistry 1 (1962) 166–171.

[89] R.V. Tomlinson, C.E. Ballou, Complete characteriza-tion of myo-inositol polyphoshpates from beef brainphosphoinositides, J. Biol. Chem. 236 (1961) 1902–1906.

[90] D. Kostrewa, F. Leitch, A.G. D’Arcy, C. Broger, D.Mitchell, A.P.G.M. vanLoon, Crystal structure of phy-tase from Aspergillus ficuum at 2.5 A, resolution, NatureStruct. Biol. 4 (1997) 185–190.

[91] J.J. Scott, F.A. Loewus, A calcium-activated phytasefrom pollen of Lilium longiflorum, Plant Physiol. 82(1986) 333–335.

[92] L. Barrietos, J.J. Scott, P.P.N. Murthy, Specificity ofhydrolysis of phytic acid by alkaline phytase from lilypollen, Plant Physiol. 106 (1994) 1489–1495.

[93] P.T. Hawkins, D.R. Poyner, T.R. Jackson, A.J.Letcher, D.A. Lander, R.F. Irvine, Inhibition of iron-catalysed hydroxyl radical formation by inositolpolyphosphates: a possible physiological function formyo-inositol hexakisphosphate, Biochem. J. 294 (1993)929–934.

[94] I.D. Spiers, S. Freeman, D.R. Poyner, C.H. Schwalbe,The first synthesis and iron binding studies of thenatural product, myo-inositol 1,2,3-trisphosphate, Te-trahedron Lett. 36 (1995) 2125–2128.

[95] B.Q. Phillipy, E. Graf, Antioxidant functions of inositol1,2,3-trisphosphate and inositol 1,2,3,6-tetrakisphos-phate, Free Radical Biol. Med. 22 (1997) 939–946.

[96] C.-F. Huang, S.M. Voglmaier, M.E. Bembenek, A.Saiardi, S.H. Snyder, Identification and purification ofdiphosphoinositol pentakisphosphate kinase, which syn-thesizes the inositol pyrophosphate bis(diphos-pho)inositol tetrakisphosphate, Biochemistry 17 (1998)14998–15005.

[97] L.R. Stephens, T. Radenberg, U. Thiel, G. Vogel, K.-H. Khoo, A. Dell, T.R. Jackson, P.T. Kawkins, G.W.Mayr, The detection, purification, structural characteri-zation and metabolism of diphosphoinositol diphospho-inoistol pentakisphosphate(s) and bisdiphosphoinositoltetrakisphosphate(s), J. Biol. Chem. 268 (1993) 4009–4015.

[98] T. Munnik, R.F. Irvine, A. Musgrave, Phospholipidsignalling in plants, Biochim. Biophys. Acta 1389 (1998)222–272.

[99] G.G. Cote, R.C. Crain, Why do plants have phospho-inositides?, BioEssays 16 (1994) 39–46.

[100] S. Carstensen, G. Pliska-Matyshak, N. Bhuvara-hamurthy, K.M. Robbins, P.P.N. Murthy, Biosynthesisand localization of phosphatidylinositol in barleyaleurone cells, Lipids 34 (1999) 67–73.

[101] Y. Pak, Y. Hong, S. Kim, T. Piccariello, R.V. Farese, J.Larner, In vivo chiro-inositol metabolism in the rat: Adefect in chiro-inositol synthesis from myo-inositol andan increased incorporation of chiro-[3H]inositol intophospholipid in the Goto-Kakizaki (G.K.) rat, Mol.Cells 8 (1998) 301–309.

[102] Y. Hong, Y. Pak, Identification of chiro-inositol-con-taining phospholipids and changes in their metabolismupon salt stress in soybean seedlings, Phytochemistry 51(1999) 861–866.

[103] A. Lykidis, P.D. Jackson, C.O. Rock, S. Jackowski,The role of CDP-diacylglycerol synthase and phos-phatidyl synthase activity levels in the regulation ofcellular phosphatidylinositol content, J. Biol. Chem. 272(1997) 33402–33409.

[104] M.R. Low, A.R. Saltiel, Structural and functional rolesof glycosylphosphatidylinositol in membranes, Science239 (1988) 268–275.

[105] M.G. Low, The glycosyl-phosphatidylinositol anchor ofmembrane proteins, Biochim. Biophys. Acta 988 (1989)427–454.

[106] A.R. Saltiel, Structural and functional roles of glyco-sylphosphoinositides, in: B.B. Biswas, S. Biswas (Eds.),Inositol Phosphates, Phosphoinositides, and SignalTransduction, Subcellular Biochemistry series, vol. 26,Plenum Press, New York, 1996, pp. 165–185.

[107] M. Kunze, J. Riedel, U. Lange, R. Hurwitz, R. Tis-chner, Evidence for the presence of GPI-anchored PM-NR in leaves of Beta 6ulgaris and from PM-NR inbarley leaves, Plant Physiol. Biochem. 35 (1997) 507–512.

[108] C. Stohr, F. Schuler, R. Tischner, Glycosyl-phos-phatidylinositol-anchored proteins exist in the plasmamembranes of Chlorella Saccharophila (Kruger) Nad-son: Plasma-membrane-bound nitrate reductase as anexample, Planta 196 (1995) 284–287.

[109] N. Morita, H. Nakazato, H. Okuyama, Y. Kim, G.A.Thompson, Evidence for a glycosylinositolphospho-lipid-anchored alkaline phosphatase in the aquatic plantSpirodela oligorrhiza, Biochem. Biophys. Acta 1290(1996) 3–62.

[110] D.K. Perry, Y.A. Hannun, The role of ceramide in cellsignaling, Biochim. Biohys. Acta 1436 (1998) 233–243.

[111] R.N. Trethewey, A.J. Krotzky, L. Willmitzer,Metabolic profiling: a Rosetta Stone for Genomics?,Curr. Opinion Plant Physiol. 2 (1999) 83–85.

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