Water Flow in the Roots of Crop Species: The Influence of ... · Structure, Aquaporin Activity, and Waterlogging ... 7. Waterlogging 171 7.1. Effect on O 2 in the rhizosphere 171

C H A P T E R T H R E E

Water Flow in the Roots of Crop

Species: The Influence of Root

Structure, Aquaporin Activity,

and Waterlogging

H. Bramley,1,* D. W. Turner,† S. D. Tyerman,* and N. C. Turner‡

Contents

1. Introduction 134

2. Water Movement Through the Plant 135

2.1. Driving forces 135

2.2. Hydraulic conductance 137

2.3. Hydraulic conductivity of roots (Lpr) 138

3. Root Characteristics and Water Flow 140

3.1. Factors that influence root growth and water uptake 140

3.2. Root anatomy 141

4. Changes in Lpr 146

5. Plant Aquaporins (AQPS) 147

5.1. AQP structure 148

5.2. AQP selectivity 150

5.3. Control of water permeability 152

6. The Role of AQPs in Root Water Transport 167

6.1. Inhibition studies 167

6.2. Expression and transformation studies 169

6.3. The contribution of AQPs to radial water flow 170

7. Waterlogging 171

7.1. Effect on O2 in the rhizosphere 171

7.2. Effect on root growth 172

7.3. Effect on water use 174

Advances in Agronomy, Volume 96 # 2007 Elsevier Inc.

ISSN 0065-2113, DOI: 10.1016/S0065-2113(07)96002-2 All rights reserved.

* Wine and Horticulture, Faculty of Agriculture, Food and Wine, The University of Adelaide (WaiteCampus), Plant Research Centre, PMB 1, Glen Osmond, South Australia 5064, Australia

{School of Plant Biology, Faculty of Natural and Agricultural Sciences, The University of Western Australia,Crawley, Western Australia 6009, Australia

{Centre for Legumes in Mediterranean Agriculture, The University of Western Australia, Crawley,Western Australia 6009, Australia

1Present address: Department of Renewable Resources, 444 Earth Sciences Building, University of Alberta,Edmonton, Alberta T6G 2E3, Canada

133

7.4. Anoxia and AQP activity 175

8. Conclusion 180

Acknowledgments 181

References 182

The hydraulic properties of plant roots depend on the morphology and anatomy

of the root system, the length of the absorbing region and the influence of

aquaporins (AQPs). These features change during development and in response

to environmental stimuli, and alter the hydraulic conductivity of the root system

(Lpr). AQPs are proteins that form water selective channels to facilitate water

flow across membranes. A large proportion of AQP isoforms are predominantly

expressed in roots and their localization indicates a putative role in the trans-

port of water across the root. AQP activity can finely regulate the rate of water

flow across the root by changes in abundance and opening/closing the water

channels. Since water will flow by the pathway of least resistance, AQPs will

only influence radial water flow if the hydraulic conductivity of the apoplast is

relatively less than that of the cell-to-cell pathway. There is growing evidence

that AQPs influence water flow through the roots of some, but not all, species.

Waterlogging is a significant environmental constraint to crop growth, but its

influence on Lpr is poorly understood. Depending on the tolerance of the

species, waterlogging through oxygen deficiency reduces root growth and

tends to reduce Lpr. Oxygen deficiency can directly or indirectly close AQPs or

alter their abundance. Changes in AQP activity may be the key component which

ultimately influences water transport through waterlogged roots.

1. Introduction

Aquaporins (AQPs) are proteins that form channels to facilitate thetransport of water across biological membranes. By altering their abundanceand/or opening and closing the channel AQPs can control the rate of waterflow into and out of cells and intracellular compartments. Since water is afundamental requirement for most life processes, there has been a prodi-gious amount of research into AQP-facilitated water transport since the firstAQP was discovered in the early 1990s. AQPs have now been found toexist in the membranes of almost all organisms, with the largest number ofAQP genes expressed in plants.

The sedentary lifestyle of plants presents a fundamental challenge in theuptake and transport of water, to meet the demands of transpiration andgrowing tissues. AQPs have the potential to mediate not only the rate offlow across membranes, but also through tissues and organs, and to provideregulation that may minimize adverse effects during abiotic perturbations.It is now widely accepted that water flow through the roots of plants is

134 H. Bramley et al.

regulated by AQP activity ( Javot and Maurel, 2002), but the majority ofthe current research tends to overlook the rudimentary features that makeplant roots successful as absorbing organs. The hydraulic properties of rootsare dependent not only on AQP activity, but also on the anatomy andmorphology of the root system, as well as the length of the absorbingregions. These features vary between species, depend on the physicalcharacteristics of the soil, and can change in response to changes in theenvironment. The nature of these changes in root structure can alter thehydraulic conductivity of the pathways for water flow through roots, eitherto conserve water during adverse conditions or function in some other way,for example, to increase the transport of gases during waterlogging. AQPactivity also varies between species and can be regulated by development,time of day, and in response to abiotic perturbation. Water flow throughroots is therefore a multifaceted process where a number of variables controlthe mechanisms.

There has been very little research into the effects of waterlogging and/orO2 deficiency on water transport through roots, despite waterlogging being asignificant constraint to crop growth. The influence of waterlogging on waterflow through roots appears to be an enigma, as unlike drought and salinitythere is an abundance of freely available water and yet root water transporttends to be reduced. Despite the effects of waterlogging on root watertransport being known for more than a century, there has been no adequateexplanation for this phenomenon. A significant discovery linked the closure ofAQPswith cytoplasmic acidosis,which occurswhen cells respire anaerobicallyduring anoxic conditions (Tournaire-Roux et al., 2003). This mechanism ofreducingmembrane permeability may be the mechanism that reduces the rateof water flow through roots, when roots are submerged during waterloggingand the O2 concentration in the rhizosphere declines.

This chapter begins with an introduction into the processes drivingwater flow and the definitions used to describe water fluxes throughroots. The review then details some of the physical characteristics thatinfluence water transport through roots and discusses how AQPs may beinvolved in regulating water flow through the radial pathway. Finally, theeffects of waterlogging on root growth and water flow through roots arereviewed. The response of AQPs to O2 deficiency is reviewed and relatedto their potential control of water flow through waterlogged roots.

2. Water Movement Through the Plant

2.1. Driving forces

Water movement in a plant is driven by gradients in water potential (Boyer,1985; Passioura, 1982). Water potential (C) is a measure of the free energyassociated with water and is expressed in units of pressure, usually

Water Flow Through Roots 135

megapascals (MPa). The net water potential of a plant is composed of twoprincipal components, hydrostatic (P) and osmotic (p) pressure:

C ¼ P � p ð1ÞP is the result of internal or external pressures, such as the tension generatedby transpiration. In cells, P includes turgor, which is an internally generatedpositive pressure, due to the rigidity of plant cell walls (Tyree and Jarvis,1982). p depends on the concentration of solutes in solution. A gravitationalcomponent is sometimes included in Eq. (1), but only for tall trees (Boyer,1985; Passioura, 1982).

Water passively moves down a gradient in total C, from high to lowenergy, until reaching equilibrium. Hence, gradients in C can drive watertransport through the soil-plant-atmosphere continuum (SPAC), and bymanipulating its C through transpiration or the accumulation of salts, aplant can control the process, within limits.

Water movement through the SPAC occurs in two phases, in the liquidphase by bulk flow through the soil and plant, and in the gas phase throughthe stomatal region (Baker, 1989). The traditional theory describing theascent of water through the SPAC is the cohesion-tension (CT) theory,although the theory is still vigorously challenged and debated (Steudle, 2001;Zimmermann et al., 1993, 2004). Transpiration during the day generateshydrostatic pressure gradients to draw water into the roots and through thexylem of the plant. This hydrostatic pressure gradient is created by the surfacetension that develops at the air–water interface in leaves and is transmitted asa negative pressure throughout the water column, where it lowersC of theroots below the soilC (Tyree, 1997). The negative pressure is equivalent to atension or pulling force, drawing water upward (Tyree, 1997). The tensionwithin the xylem increases as the soil water decreases and/or as the transpi-ration rate increases (Tyree and Sperry, 1988). By controlling the size of thestomatal apertures, the plant can regulate transpiration, and therefore P, toavoid an excessive amount of cavitation. However, when stomata are closed,photosynthesis is also inhibited, creating a compromise betweenCO2 uptakeand water loss.

The accumulation of solutes across semipermeablemembranes also estab-lishes a gradient in C that induces water uptake (Oertli, 1991). An osmoticgradient generates root pressure, causing xylem sap to exude from a cut shootor detopped root system (reviewed by Zholkevich, 1991). The endodermisprobably forms a semipermeable barrier, preventing the net efflux of solutesfrom the stele. Solutes secreted into the xylem lower C and induce wateruptake across the root. An upward flow of solution results as more water isdrawn into the xylem.

Osmotic gradients are only important in driving water transport whentranspiration is low (Kramer, 1983). When transpiration increases, the


increasing mass flow of water dilutes the solutes in the xylem sap until theosmotic component is negligible and the uptake of water is predominantlycontrolled by the hydrostatic pressure within the xylem (Section 2.3).Transpiration can produce a much steeper gradient in C, from soil toroots, than an osmotic mechanism, as reviewed by Kramer (1983). Evenso, water uptake may be limited when a plant experiences water deficitthrough drying soil or saline conditions because the plant cannot lower itsCsufficiently to create a gradient between the soil and roots.

2.2. Hydraulic conductance

The flux of water (rate of water flow) through a plant depends not only onthe size of the driving forces, but also on the conductance (reciprocal ofresistance) of the pathways, through which water flows (Boyer, 1985):

Q ¼ LðDP � sDpÞ ð2Þ

where Q is the water flux or volume flow rate (m3 s�1), L the hydraulicconductance (m3 s�1 MPa�1), s the reflection coefficient, andDP andDp thehydrostatic and osmotic pressure differences (MPa), respectively. s is a unitlessparameter, relating the interaction between water and solute crossing a mem-brane. The value of s depends on the particular solute, with a value of 1 for aperfect osmometer and 0 when the membrane does not reflect the soluterelative to water. If s ¼ 1, then Eq. (2) reduces to (Boyer, 1985):

Q ¼ LðDCÞ ¼ ðDCÞR

ð3Þ

where R is the apparent hydraulic resistance (MPa s m�3) and hence, Eq. (3)is a simple analogy to Ohm’s law (van den Honert, 1948).

A common error in plant water relations is the interchangeable use ofhydraulic conductance and conductivity (Lp). Equation (4) shows therelationship between the two parameters. L is a measure of the ability ofan entity to conduct water, independent of the entity’s dimensions, whereasLp is a property of an entity with specified dimensions, usually surface area(A).

L ¼ Lp�A ð4ÞThe SPAC has been described as a system of hydraulic resistors arranged inseries (van den Honert, 1948). Plants can vary the resistance (and conduc-tance) of the pathways to maintain the water balance of the shoot (Steudle,2000). When water exists in the vapor phase, the greatest resistance is thestomatal aperture. However, in the liquid phase, the root system constitutes


a highly significant and important resistance to overall flow of water in theplant (Steudle, 2000).

2.3. Hydraulic conductivity of roots (Lpr)

The surface area limits water uptake by the root (Steudle, 2001; Tyree,2003). Therefore, water transport parameters are usually normalized perunit surface area and revision of Eq. (2) incorporates this:

Jv ¼ LprðDP � srDpÞ ð5Þ

where Jv is the volume flow, per unit area (m3 m�2 s�1 or m s�1), Lpr thehydraulic conductivity of the root (m s�1 MPa�1), and sr the apparent rootreflection coefficient (Passioura, 1988). The relative influences of DP andDp depend on the rate of salt secretion into the xylem and the rate of waterflow (Weatherley, 1982). When DP is zero, water flow is driven by theosmotic gradient, but the xylem sap becomes diluted when water flowincreases through transpiration, and Dp becomes negligible (Kramer, 1983).

The majority of studies have used detopped root systems to measure Jv,where the exudation of sap from the cut stump is collected under rootpressure, or water flow is induced either by applying suction to the cutstump or externally pressurizing the root system (Fiscus, 1975; House andFindlay, 1966; Nobel et al., 1990). In the root pressure exudation technique,the driving force is osmotic and Dp is determined by measuring the osmoticpressure of the exudate and the medium bathing the roots.

In the external pressure technique, the root system is sealed in a pressurechamber with the cut stem protruding through the lid of the chamber. Theroot system is pressurized and Jv determined at different pressure increments.Plotting Jv against DP typically produces a curvilinear relationship (Passioura,1984, 1988), due to the osmotic component dominating flow at low flow rates(Dalton et al., 1975; Fiscus, 1975). At high flow rates, the solutes are so dilutedthat Jv(DP) is inherently linear and the slope givesLpr [Eq. (5)]. If the linear partof the Jv(DP) relationship is extrapolated to the x-axis, the intercept shouldequal the osmotic pressure of the external medium (po), but invariably itexceeds po (Passioura, 1984). This anomaly has not been satisfactorilyexplained (Passioura, 1984), but the offset depends on the species and canvary diurnally or with abiotic perturbations (Boursiac et al., 2005; Emery andSalon, 2002; Munns and Passioura, 1984; Murphy, 2003; Passioura andMunns, 1984; Rieger and Litvin, 1999).

The root and cell pressure probes have been developed to measure waterflow through individual roots and across cell membranes (Steudle, 1993;Tomos and Leigh, 1999). In summary, the root pressure probe (RPP)measures root pressure and water flow is induced either by applying


hydrostatic pressure (with the aid of the probe) to the root, or by changingthe osmotic pressure of the bathing medium. The rate of water flow acrossthe root, and subsequently, Lpr is calculated from the rate of transientrelaxations in root pressure (Steudle, 1993).

The cell pressure probe (CPP) measures turgor pressure via the tip of amicrocapillary introduced into an intact cell. Analogous to the RPP, waterflow is induced by applying hydrostatic pressure, with the aid of the probe(Steudle, 1993). If the dimensions of the cell are known, the CPP can beused to determine the volumetric elastic modulus (e) and hydraulic con-ductivity (Lpc) of the cell. Osmotic flows can also be induced, but due to theuncertainty of the effects of unstirred layers for cells located within tissue,the values of Lpc may be erroneous.

The CPP andRPP are appealing because they provide real-time measure-ments that are useful for estimating the location of the principal resistances towater transport through roots. Measurements of cell turgor have revealedgradients in C across a radial profile in the roots of some species (Pritchardet al., 1989; Zimmermann et al., 1992). Combining results from CPP andRPP measurements on roots of the same species can potentially identify themain radial pathway for water transport across the root (Section 3.2.2). Forexample, Lpc of epidermal and cortical cells was much greater than Lpr ofHordeum distichon and Phaseolus coccineus roots, indicating that water flows viathe cell-to-cell pathway (Steudle and Brinckmann, 1989; Steudle and Jeschke,1983). In comparison, analogous measurements on maize (Zea mays) rootsrevealed a predominantly apoplastic flow (Steudle et al., 1987). Comparing themeasured values of Lpr and Lpc for each cortical cell layer indicated that radialwater flow throughwheat (Triticum aestivum) roots occurs by a combination ofthe parallel pathways, but radial water flow in the roots of narrow-leafed lupin(Lupinus angustifolius) and yellow lupin (L. luteus) appears to be predominantlyapoplastic (Bramley, 2006).

2.3.1. Transport modelsThe root has been modeled as a system containing either two or threecompartments, a series of membranes or as a perfect osmometer, to explainthe processes of water flow across plant roots (Dalton et al., 1975; Fiscus,1975; Pickard, 2003; Tyree et al., 1994). However, there has been consid-erable debate about whether Lpr of roots was dependent on Jv (reviewed by(Kramer, 1983; Passioura, 1982). Before the models of Fiscus (1975) andDalton et al. (1975), the nonlinear relationship between flow and drivingforce was interpreted as Lpr increasing with flow rate, so water flux wasconsidered to be independent of DC across the root (reviewed byWeatherley, 1982). The landmark papers of Passioura and Munns (Munnsand Passioura, 1984; Passioura and Munns, 1984) demonstrated a linearrelationship between driving force and flow. The slope of the relationship


varied with time and treatment, indicating that the apparent hydraulicconductance was truly variable.

Steudle (1994) extended the two compartment models and developedthe Composite Transport model. The model incorporates the compositenature of water flow across different tissues, the parallel radial pathways,the hydrostatic and osmotic driving forces, variable Lpr, and also accountsfor sr < 1 (Steudle and Peterson, 1998). The model explains the variabilityin Lpr in relation to the nature and intensity of the driving forces (Steudleand Peterson, 1998). Osmotic forces will primarily drive water flowthrough the cell-to-cell pathway, as s�1 for cell membranes. Since thereare no membranes in the apoplastic pathway, the apoplastic s�0. Thus,osmotic gradients will drive very little water transport in the apoplast. Whenthe driving force is primarily hydrostatic (e.g., generated by transpiration),water can flow via a combination of apoplastic and cell-to-cell pathways.

3. Root Characteristics and Water Flow

3.1. Factors that influence root growth and water uptake

External factors can influence water uptake by the roots, such as interactionsbetween the roots and the soil, and the distribution of roots within the soilprofile (Passioura, 1988). Close contact with the soil is imperative for rootsto maintain hydraulic continuity at the soil–root interface and minimize theinterfacial resistance to flow. Poor hydraulic contact necessitates a large dropin water potential across the interface to induce water uptake (Passioura,1988). Roots growing through large soil pores may have poor hydrauliccontact with the soil, but there is also evidence that some roots shrink whenplant C is low, so the root–soil contact may be reduced (Huck et al., 1970;Palta et al., 1987). The growth of root hairs or the exudation of compoundsthat adhere to soil particles may assist roots in maintaining an intimateconnection with the soil.

The distribution of roots in the soil profile depends on soil texture andstructure, and the type of inherited root system (Kramer, 1983). Roots tendto grow through preexisting soil pores, toward regions with nutrients andwater, and avoid unfavorable regions, so the distribution is generallynot uniform. Roots of many crop species can penetrate to depths of severalmeters in well-aerated, deep, soft soils (Kramer, 1983). In addition, toler-ance to the range of abiotic limitations encountered by roots in a growingseason may determine root growth behavior.

There are two main types of root system. Monocots develop a fibrousroot system with initial roots (3–5 axes for wheat) emerging from the seed(seminal roots) and subsequent roots emerging from the basal nodes of thestem (nodal roots, also called adventitious roots) (Greacen et al., 1976).In comparison, the radicle of eudicots (the first root to emerge from the


seed) can develop into the dominant root of a taproot system that canbecome extensively branched. Although monocot and eudicot root systemscan extend to similar depths in the field, the total root length density (rootlength/soil volume) of monocots is often much greater. For example, theroot length density of wheat is two- to tenfold larger than narrow-leafedlupin in Western Australian soils (Dracup et al., 1993; Gallardo et al., 1996;Gregory and Eastham, 1996; Hamblin and Tennant, 1987). Despite thesedifferences in root length, the roots of eudicotyledon species tend to have ahigher specific rate of water uptake than cereals (Bremner et al., 1986;Hamblin and Tennant, 1987; Mason et al., 1983). Moreover, these greaterrates of water uptake appear to be due to a greater hydraulic conductivity.For example, Lpr of lupin root systems is at least twofold greater than Lpr ofwheat root systems (Bramley, 2006; Gallardo et al., 1996).

3.2. Root anatomy

Kramer (1983) stated that the ‘‘effectiveness of roots as absorbing organs’’depends on anatomical and morphological features. The dynamics of rootwater permeability in relation to these features were succinctly summarizedby Moreshet and Huck (1991). The root apex typically has high axial andradial resistances to water flow compared with the remainder of the rootwith its developed xylem (Steudle, 2001). Water and nutrient absorptiongenerally commences >10 mm behind the tip, which coincides with theroot hair zone. The length of the absorbing region depends on the species,but may change with transpirational demand and during adverse conditions(reviewed by Kramer, 1983). Lpr of wheat roots decreases with distancefrom the root tip, indicating that water absorption occurs preferentially inthe apical region (Bramley, 2006; Jones et al., 1988). In addition, waterabsorption by several or all of the individual roots may contribute to Lpr ofthe whole root system (Bramley, 2006). Cereal roots apparently havemaximum water absorption within a region <100 mm from the root tip(Greacen et al., 1976; Sanderson, 1983). There is also evidence that individ-ual roots are able to vary their hydraulic conductivity. Vysotskaya et al.(2004a,b) excised four of the seminal roots of durum wheat (Triticumdurum), and this increased Lpr of the remaining root, so that the watersupply to the shoot was maintained. In comparison, lupin roots absorbwater more evenly along the length of the taproot as Lpr is constant withlength (Bramley, 2006). Using a novel CAT-scanning and microelectrodetechnique, Hamza and Aylmore (1992a) observed uniform water absorptionalong the length of roots of 17-day-old narrow-leafed lupin plants, incomparison with radish (Raphanus sativus) plants that extracted more watercloser to the soil surface. Lupins had lower potential difference betweenleaves and root surface (i.e., smaller DC, which equates to a smaller drivingforce) and higher water flow rates than radish, indicating that lupins hadlower plant resistances (Hamza and Aylmore, 1992b).


This disparity in root water uptake and Lpr were initially thought to bedue to differences in axial and radial anatomy (Gallardo et al., 1996; Hamblinand Tennant, 1987; Hamza and Aylmore, 1992a), but this was proposedbefore AQPs were discovered. Nevertheless, differences in anatomy caninfluence the hydraulic conductivity of the pathways for water flow.

The root has been described as a porous pipe or leaky cable, where the ratioof the radial (across the pipe wall) and axial (through the length of the pipe)resistances determines the resistance of the whole root and the distribution ofwater uptake (Landsberg and Fowkes, 1978; Zwieniecki et al., 2003). In theradial pathway, water taken up by the root has to traverse living tissue beforereaching the lumen of the xylem vessels. In the axial plane, water flow occursthrough the xylem vessels and tracheids.

3.2.1. Axial pathwayMature xylem vessels are dead cells that form a continuum of tube-likestructures, separated by thin perforatedwalls, so that longitudinal flow throughthe vessels is analogous to flow through a conduit or pipe (Zwieniecki et al.,2003). The driving force for longitudinal flow through the vessels will behydrostatic, unless membranes (e.g., living cells) interrupt the pathway. If theperforated walls provide negligible resistance, the radius of the vessels (to thefourth power) determines the rate of water flow through the xylem contin-uum, according to the Poiseuille-Hagen equation (Steudle and Peterson,1998). The development of the vasculature is different between monocotsand eudicots. For example, the stele of wheat contains one central pittedmetaxylem vessel, which increases in diameter away from the root tip (40-to 100-mm diameter), and seven or eight xylem strands containing one smallmetaxylem vessel (10-mm diameter). Consequently axial conductance doesnot appear to change much along the length of the root (Bramley, 2006).Within the stele of lupins, the vasculature develops in a diarch pattern, withbundles of metaxylem vessels up to 100 mm in diameter (Hamblin andTennant, 1987). The abundance and diameter of vessels increase with distancefrom the root tip, which result in axial conductance also increasing in a similarpattern (Bramley, 2006).

The axial conductance may be associated with Lpr, probably reflecting thecapacity of the axial pathway in relation to all water absorption distal to anypoint. The axial conductance may be several orders of magnitude greater thanthe total hydraulic conductance of a root and hence the radial pathway createsthe greatest constraint on water flow through the root (reviewed by Steudleand Peterson, 1998).

3.2.2. Radial pathwayWater taken up by the root has to cross a series of concentric cell layers: theepidermis, cortex, endodermis, and stele, before reaching the lumen ofthe xylem vessels (Fig. 1).


The epidermis consists of a single layer of elongated, tightly packed cells.Epidermal cell walls are generally thin, but may contain deposits of suberinor cutin (Kramer, 1983; Moreshet and Huck, 1991). Epidermal walls of themajority of 181 species surveyed appeared autoflourescent when viewedunder ultraviolet light, indicating the presence of suberin or lignin(Perumalla et al., 1990). Root hairs arise from epidermal cells, and theirabundance and longevity depend on the plant species and environment.A structurally different subepidermal layer may exist, called the hypodermis(Perumalla et al., 1990). In some species, the hypodermal walls contain aradial strip of suberin (Casparian bands) or suberized secondary walls and thelayer is called an exodermis (Enstone and Peterson, 1997, 1998; Perumallaand Peterson, 1986; Perumalla et al., 1990). Suberin is composed of hydro-phobic, fatty compounds, believed to act like a waterproofing agent(Nawrath, 2003; Zeier and Schreiber, 1998; Zeier et al., 1999). The pres-ence of a suberized exodermis can change with root maturity and environ-mental conditions. For example, in maize roots, aeroponic culture (mistculture) induces the development of an exodermis (Hose et al., 2000;Zimmermann et al., 2000). In comparison, the roots of lupin species donot form a hypodermis or exodermis, at least up to 300 mm from the tipwhen grown in hydroponic or sand culture (Bramley, 2006; Hartung et al.,2002; Perumalla et al., 1990) and cannot be stimulated by aeroponics(Hartung et al., 2002). A suberized exodermis does not form in membersof the Tritaceae family (Greacen et al., 1976; Perumalla et al., 1990) possiblybecause the cortex eventually deteriorates.

The cortex consists of cell layers, the number of which varies dependingon species, root development, and environment. The cortex of cereal rootsgenerally consists of 6–8 cell layers, but this shrivels with maturity (Greacenet al., 1976; Hamblin and Tennant, 1987). The cortex also shrinks when thewater potential of the root decreases, which implies it is not buffered against

B

epC

en

A

epC

en

X

Figure 1 Transverse sectionsof (A)wheatand(B)narrow-leafed lupin roots,20^40 mmfromthe root tip, showing the radial cell layers that water must cross before reaching thelumen of the xylem vessels. For wheat and lupins, water must cross the epidermal (ep),cortical (C), endodermal (en), and stelarcell layers. Scale bars represent 50 mm.


large changes in C in the xylem. Hence, the greatest resistance to radialwater flow may occur in the epidermis (Passioura, 1988). Cortical cell wallsare thin and usually contain no suberin. The cubic or hexagonal arrange-ment of cells determines the shape of the intercellular spaces between cells,which influences the air-filled root porosity and the size of the apoplast.

The endodermis is a single cell layer that forms a sheath around the stele(Moreshet and Huck, 1991). The cell walls of the endodermis may containsuberin and thicken to form a similar structure to the exodermis (Perumallaand Peterson, 1986; Zeier and Schreiber, 1998; Zeier et al., 1999). Duringroot growth, the endodermis develops in three stages: (1) appearance ofCasparian strip; (2) continuous layer of suberin (suberin lamellae) coveringthe protoplast, between plasmalemma and cell wall; and (3) inner tangentialwalls thicken, and a layer of cellulose is deposited over the suberin lamellae(Ma and Peterson, 2003). The location of the different endodermal stagesalong the root axis depends on species, but appears to mature earlier incereals compared with eudicots (Bramley, 2006; Greacen et al., 1976;Hartung et al., 2002; Perumalla et al., 1990; Sanderson, 1983). Water deficitmay also promote earlier development of the endodermis (Enstone et al.,2003). The endodermis may also contain unsuberized passage cells, oppositexylem poles (Esau, 1977).

The pericycle, which is the layer from which lateral roots arise, boundsthe stele. The stele contains small, tightly packed parenchyma cells and thevascular tissue (xylem and phloem). Primary xylem forms a core in the rootthat develops in an exarch pattern, that is, centripetally (Esau, 1977).Protoxylem elongates and matures (deposition of secondary walls) tobecome a functioning conduit, before eventually differentiating intometaxylem.

3.2.2.1. Parallel pathways for radial water flow There are three parallelpathways for radial water flow: symplastic, transmembrane, and apoplastic(Fig. 2).

The symplastic pathway is from cell to cell, through the cytoplasmiccontinuum, via plasmodesmata (Fig. 2). Water must cross at least twomembranes, once to enter and once to exit the symplast.

The transmembrane pathway involves water flow from cell to cell(vacuole to vacuole), and the most direct pathway is by crossing cell walls,plasma membrane, cytoplasm, and tonoplast (Fig. 2). Water crossing eachcell layer in series, via the transmembrane pathway, would hence cross fourmembranes per cell (Fig. 2).

The apoplastic pathway is around protoplasts, via the cell walls andintercellular spaces (Fig. 2). Analogous to flow through the xylem, hydro-static pressure gradients drive water flow in the apoplastic pathway becauseit contains no membranes. However, the apoplastic path has small cross-sectional surface area of the radial pathway and may be interrupted at the


endodermis and exodermis (if present) by suberin, forcing water to cross atleast the plasma membrane to reach the apoplast of the stele. Suberization ofthe endodermis and exodermis is thought to block the apoplast and preventthe transport of water and ions (Enstone et al., 2003), but Steudle et al.(1993) demonstrated that the endodermis of young maize roots did notinfluence Lpr.

Water flow across the root can occur by any combination of apoplasticand cell-to-cell pathways, or one pathway may dominate. The pathwaysmay differ between species, change in different regions of the root, and alterwith different environmental conditions. Identifying the contribution ofthe different pathways is important for understanding the nature of theresistances to root water transport. Some researchers believe the cell-to-cellpathway is the main pathway for water flow, and that the epidermis poses thegreatest resistance (reviewed by Passioura, 1988). Transport through the cell-to-cell pathway, crossing membranes or via plasmodesmata, provides theopportunity for regulatory control and hence the potential to influence thesize of the resistance, without necessitating anatomical changes. Moreover,

X

V

Plasma membrane PlasmodesmataTonoplast Cell wall

CbCb

Epidermis Exodermis Cortex Endodermis Stele

Apoplastic path

Symplastic path

Transmembrane pathCell-to-cell path

Figure 2 Pathways of radial water flow, from the root surface to the lumina of xylemvessels (X).The apoplastic pathway (cell walls and intercellular spaces) may be blockedat the endodermis and exodermis (if present) by the Casparian strip (Cb).The symplas-tic path is within the cytoplasm, through plasmodesmata. Flow in the transmembranepath crosses the plasmamembrane and vacuole (V).


the discovery of AQPs is changing the traditional view that water transportacross membranes has a high invariant resistance to water flow (Tyerman et al.,1999).

The apoplast is often considered to be the dominant pathway, despitethe fact that it only constitutes �5% of total root volume and an evensmaller fraction of the cross-sectional surface area. If the apoplast were themain pathway, the plant would have little control of water flow throughthe root, unless the endodermis forms a tight barrier and therefore, plays acentral regulatory role.

If the radial conductance is indeed the rate-limiting factor in watertransport through roots, then theoretically, thicker roots should have asmaller Lpr than thin roots because the path length for flow is longer.However, anatomy and AQP activity (Section 6.3) influence the resistivityof the radial pathways to flow, which can counteract the significance of thepath length. Rieger and Litvin (1999) found that root diameter was nega-tively correlated with Lpr in five species with contrasting root anatomy. Inaddition, Rieger and Litvin (1999) found that drought stimulated suberiza-tion and other anatomical changes that reduced Lpr. The development of anexodermis in maize roots decreases the radial hydraulic conductivity (Hoseet al., 2000; Zimmermann et al., 2000). Conversely, Barrowclough et al.(2000) discovered that in developing onion (Allium cepa) roots, the greatestvalues of radial hydraulic conductivity were correlated with the presence ofan exodermis. Huang and Eissenstat (2000) also found that structural differ-ences in the radial pathway were the primary factor determining Lpr of rootsof citrus rootstocks.

4. Changes in Lpr

Lpr for individual roots and whole root systems typically rangesbetween 10–8 and 10–7 m s�1 MPa�1, for herbaceous species, reflectingdifferences in root structure and experimental techniques (Table 6 ofSteudle, 1989; Table 2 of Rieger and Litvin, 1999). For any given species,Lpr can alter with development, environment, and other regulatory controls.

During the day, water flux through the plant varies with time, usuallyfollowing the fluctuation in transpiration. A number of studies have found Lprto vary diurnally in excised roots of wheat, maize, Lotus japonicus, castor oil(Ricinus communis), white lupin (L. albus), and sunflower (Helianthus annuus)(Carvajal et al., 1996; Clarkson et al., 2000; Else et al., 2001; Everard andDrew,1987, 1989; Henzler et al., 1999; Passioura and Munns, 1984).

Exogenous addition of phytohormones to the root-bathing mediumtends to influence root water transport, although the responses to abscisicacid (ABA) treatment are contradictory. The hydraulic conductivity ofmaize, sunflower, and common bean (Phaseolus vulgaris) roots increased in


response to exogenous ABA (Aroca et al., 2006; Freundl et al., 2000; Hoseet al., 2000; Quintero et al., 1999). ABA also increased Lpc of maize rootcortical cells, but the effect was only transient with Lpc returning to controllevels after 2 h (Hose et al., 2000). Conversely, ABA did not influence waterflux from detopped Populus tremuloides roots, but reduced Lpr of P. coccineusroots (Fiscus, 1981; Wan and Zwiazek, 1999).

Abiotic factors also affect hydraulic conductivity. Gradual soil drying oraddition of external osmoticants (e.g., NaCl, polyethylene glycol) decreasedroot water transport by 27–100% in long- and short-term experiments(Aroca et al., 2006; Azaizeh and Steudle, 1991; Carvajal et al., 1999, 2000;Joly, 1989; Liu et al., 2006; Lu and Neumann, 1999; Martre et al., 2001;Munns and Passioura, 1984; Nobel et al., 1990; Rieger and Litvin, 1999).Other factors that can reduce Lpr include O2 deficiency (Section 7.3),nutrient deficiencies, chilling, and high concentrations of aluminium orother toxicants (Aroca et al., 2005; Carvajal et al., 1996; Clarkson et al.,2000; Gunse et al., 1997; Kamaluddin and Zwiazek, 2002a, 2003; Lee et al.,2004). However, the results are not consistent, and for example, starving theroots of potassium more than doubled Lpr of rice (Oryza sativa) roots, whichwas correlated with an increase in expression of AQP genes (Liu et al.,2006). The response of Jv and Lpr is frequently rapid, occurring withinminutes and when the treatment is removed the recovery in Lpr is often justas rapid (Carvajal et al., 1996; Gaspar et al., 2001; Kamaluddin and Zwiazek,2002a, 2003). The magnitude of the effect on Lpr appears to depend on thenature of the driving force with salinity and chilling inhibiting osmoticLpr more than hydrostatic Lpr (Azaizeh and Steudle, 1991; Lee et al., 2004;Martinez-Ballesta et al., 2000).

Physical changes in root structure and anatomy occur in some species(Section 3) and these may decrease the conductance to bulk water flow, forexample deposition of suberin in hypodermal cell walls (Steudle, 2000).However, anatomical changes are slow and dependent on growth, andmay act as a survival strategy to reduce Lpr in the long term, whenenvironmental changes in the field are slow. There is no evidence whereanatomical changes provide the means to regulate Lpr diurnally, nor do theyaccount for the rapid and reversible changes in Lpr measured under labora-tory conditions. If a significant proportion of radial water flow occursthrough the cell-to-cell pathway by crossing cell membranes, Lpr may becontrolled by AQP activity.

5. Plant Aquaporins (AQPS)

Plant AQPs are ubiquitous and expressed at such high levels that theycan constitute up to 15% of total membrane protein (Johansson et al., 1996;Rivers et al., 1997). Plant AQPs are also highly diverse. The Arabidopsis


thaliana genome contains 35 sequences that are AQP homologues andat least 31 AQPs have been identified in maize and 33 in rice (Chaumontet al., 2001; Quigley et al., 2001; Sakurai et al., 2005). The high number ofAQPs alludes to their importance in plant membrane transport and possiblyplant hydraulics (The Arabidopsis Genome Initiative, 2000). Based on thesequence alignment of their amino acids, the plant AQP family is classifiedinto four subfamilies: plasma membrane intrinsic proteins (PIPs), tonoplastintrinsic proteins (TIPs), nodulin-like intrinsic proteins (NIPs), and smallbasic intrinsic proteins (SIPs). The PIPs are further subdivided into twogroups, PIP1 and PIP2. The PIP and TIP nomenclature implies that theseproteins are localized to the respective plasma and vacuolar membranes, butthey may also be localized to other membrane compartments (Barkla et al.,1999; Frangne et al., 2001; Kirch et al., 2000; Liu et al., 2003).

The diversity of plant AQPs implies different functional roles, althoughthe nature of these roles is still generally unclear. Phylogenetic analysis showsclose similarities between monocots and eudicots and the genomes ofA. thaliana, rice and maize contain similar numbers of AQP homologueswithin each subfamily/group (Table 1). Rice has two less PIP members thanA. thaliana and maize, and maize has fewer NIPs compared with the othertwo species. Although there is a high degree of homology between AQPswithin each subfamily, each species also has unique AQP members(Chaumont et al., 2001; Johanson et al., 2001; Quigley et al., 2001; Sakuraiet al., 2005).

5.1. AQP structure

In AQP proteins there are six transmembrane helices connected by fiveloops of varying lengths and the amino (NH2) and carboxy (COOH)termini are located on the cytoplasmic side of the membrane (Fig. 3).Amino acid motifs in loops C and E are highly conserved in a wide rangeof PIPs, but are not present in AQPs localized in the tonoplast, and so thesemotifs are thought to be necessary for signal recognition and targeting to therespective membrane (Barone et al., 1997).

The two longest loops, B and E, contain the signature AQP motifsasparagine-proline-alanine (NPA) that fold into the membrane as a compo-nent of the single aqueous pore (Fig. 3). The novel folding of the polypeptidecreates a pore shape that is analogous to an hourglass, with a narrow constric-tion in the center of the membrane and widens at the membrane surfaces( Jung et al., 1994; Murata et al., 2000). The obverse symmetry of the poreprovides AQPs with bi-directionality without rectification (Meinild et al.,1998; Murata et al., 2000). LoopD in PIPs is four to seven amino acid residueslonger than other AQPs, which folds under the protein on the cytosolic sideof the membrane and forms a cap occluding the cytosolic side of the pore(Tornroth-Horsefield et al., 2006, Section 5.3.3.1.1).


A cysteine residue on the third transmembrane domain (Fig. 3) has beenfound to impart mercury sensitivity to several plant AQPs and this residue ishighly conserved in PIPs and TIPs (Daniels et al., 1996; Quigley et al.,2001). Barone et al. (1997, 1998) also demonstrated that Hg 2þ induces aconformational change to the plant plasma membrane AQPs instead ofdirect occlusion of the water channel. Several PIPs and at least one TIPappear to be mercury insensitive, despite the presence of several cysteineresidues on the protein (Biela et al., 1999; Daniels et al., 1994; Krajinski et al.,2000). Despite the caveats of using this compound (Section 5.3.3.1.3), theapplication of mercury is one of the main tools used to estimate the activityof AQPs by determining its effect on reducing membrane permeability(Maurel, 1997).

Other features that are highly conserved in plant AQPs include a pH-sensitive histidine residue on Loop D (Fig. 3), present in all PIPs(Tournaire-Roux et al., 2003). Serine residues, which are putative phos-phorylation sites, are located in the first cytosolic loop of most PIPs and

Table 1 The number of genes identified within each AQP group in the genomesequences of arabidopsis, rice, and maize

AQPsubfamily/group

Arabidopsisthaliana

Orysasativa Zeamays References

PIP1 5 3 6 Chaumont et al., 2001;

Johanson et al., 2001;

Quigley et al., 2001;

Sakurai et al., 2005

PIP2 8 8 7 Chaumont et al., 2001;




TIP 10 10 11 Chaumont et al., 2001;




NIP 9 10 3 Chaumont et al., 2001;




SIP 3 2 4 Chaumont et al., 2001;





TIPs, and the C-terminal of all PIP2s and NIPs (Fig. 3) (Guenther et al.,2003; Johansson et al., 1998; Quigley et al., 2001).

In the membrane, AQPs form tetrameric structures with each monomeracting as an independent water channel (Murata et al., 2000;Walz et al., 1997).There is increasing evidence that at least some plant AQPs form hetero-tetramers through interaction with different PIP isoforms and this inter-action activates water channel function (Fetter et al., 2004; Temmeiet al., 2005). However, for functional importance to be implied, studies needto demonstrate that the relevant isoforms are coexpressed in the same tissueand cells.

5.2. AQP selectivity

AQP activity is predominantly assayed by heterologous expression of indi-vidual isoforms in Xenopus laevis oocytes. The oocytes have an intrinsicallylow permeability to water and are invariably unperturbed when exposed toa hypotonic bathing medium. After expression of exogenous AQPs theoocytes swell rapidly and burst within a few minutes. The swelling rate isused to calculate the osmotic water permeability of the membrane (Pf).Although this method does not reveal the true function of AQPs in nativemembranes, the procedure does demonstrate that many AQP homologuesincrease the rate of water flow across membranes. The activity of individual

Apoplast or vacuole

Cytosol

H2N

HOOC

A

1 2 3 4 5 6

B C D E

P

PNN

AA

C

S H

S

Figure 3 Overall topology of a plant PIP2 showing the highly conserved and impor-tant residues (circled). AQPs contain six membrane-spanning domains connected byloops Ato E. LoopDof PIPs contains four to sevenmore amino acid residues than otherAQPs and this loop folds under the protein, on the cytosolic side, to block the pore.TheNPAmotifs form the aqueous pore. Serine (S) and histidine (H) residues are involved inphosphorylation and protonation of the protein, respectively.The cysteine (C) residueon the third transmembrane domain confersmercury sensitivity in someAQPs.


AQPs varies considerably, with some isoforms having very low waterpermeability, while others are also permeable to small nonelectrolytes,such as CO2, glycerol (aquaglyceroporins), boric acid, and urea (Bielaet al., 1999; Dordas et al., 2000; Liu et al., 2003; Uehlein et al., 2003).PIP1s appear to have little or no water permeability, although whenexpressed individually they may not be activated in oocytes (Chaumontet al., 2000; Fetter et al., 2004; Li et al., 2000; Suga and Maeshima, 2004).Conversely, PIP2s have high water permeability and can increase Pf by atleast tenfold (Chaumont et al., 2000; Suga and Maeshima, 2004). NOD26from soybean (Glycine max) and its ortholog LIMP2 from L. japonicus havemoderate water permeability and only increase oocyte swelling rates bythreefold, but are also permeable to glycerol (Guenther and Roberts, 2000;Guenther et al., 2003). NtAQP1 from tobacco (Nicotiana tabacum), a PIP1ortholog, is moderately permeable to water, glycerol, and CO2 (Biela et al.,1999; Siefritz et al., 2001; Uehlein et al., 2003). TIPs appear to haveconstitutively high permeability to water and several A. thaliana TIPs alsotransport urea (Daniels et al., 1996; Liu et al., 2003). No plant AQPs thathave been functionally characterized are permeable to ions, unlike themammalian AQP6 that is permeable to anions, particularly nitrate, whenexposed to HgCl2 or when the cytosol is acidified (Hazama et al., 2002;Ikeda et al., 2002; Yasui et al., 1999). HgCl2 can cause turgor pressure ofplant cells to decrease due to a leakage of ions (Zhang and Tyerman, 1999),albeit the loss of ions is probably through ion channels.

The shape of the pore and the size of the permeating molecule confer theselectivity of the individual AQP for water or other small neutral solutes(Murata et al., 2000). However, a simple steric model does not account forthe diversity of solutes that can permeate some AQPs and not others.As Meinild et al. (1998) demonstrated, there is no connection betweenthe permeation of mammalian AQPs and the solute size. Selectivity of theprotein is more likely determined by the properties of the amino acidresidues lining the pore. Identification of the important residues for selec-tivity is progressing for prokaryotic and mammalian AQPs (Sui et al., 2001;Tajkhorshid et al., 2002), but no specific signature motifs have been identi-fied in solute-transporting plant AQPs thus far (reviewed by Santoni et al.,2000; Tyerman et al., 2002). There is also some conjecture whether thefourfold axis in the center of the AQP tetramer could function as a channel,and possibly as an ion channel, which is activated when the state of the waterchannels is altered (Chrispeels et al., 1999; Tyerman et al., 2002).

Considerable advances have been made in determining the permeabilityof different AQPs to various compounds, but knowledge of the specifics ofpermeability alone does not infer a physiological role. With new technologyand molecular techniques, research is progressing in understanding of therole of AQPs in situ.


5.3. Control of water permeability

The existence of AQPs is profoundly significant for living cells, becausethey endow the organism with the potential to control water flow(Tyerman et al., 2002). AQPs can control flow across the membrane (andtissue/organs) in two ways: (1) by changing their abundance in the mem-brane and/or (2) by changing the rate of flow through the water channel.Changing the rate of flow through the AQP may be achieved by theopening and closing of the channel (gating), or by changing the gradientof the driving force (Tyerman et al., 2002).

5.3.1. AQP abundance and patterns of expression in rootsThe expression pattern of AQPs tends to be greater in tissues and cells withhigh water permeability. In plants, the patterns of expression are complex,varying between organs and tissues and are dependent on the species. Veryfew AQP isoforms appear to be cell specific, but rather, are expressed atvarying levels in a range of cell types under developmental control or inresponse to environmental stimuli.

When focusing on roots, the literature reveals several principal features.First, a high proportion of AQP isoforms is preferentially expressed in roots(Table 2). PIP2;1 and/or PIP2;2 homologues are abundantly expressed inthe roots of most species studied (Table 2). The tissue-specific expression ofthese isoforms also reveals a putative role in the radial transport of wateracross the root with the high abundance of PIP2 homologues particularly inthe endodermis (Hachez et al., 2006; Javot et al., 2003; Suga et al., 2003).If suberization of the endodermis blocks the apoplastic pathway, then waterwould have to cross membranes in this tissue to reach the apoplast of thestele (Section 3.2.2.1). Geometrically, the endodermis also creates a highresistance to water flow, because of the decreasing surface area towardthe center of the root. A high expression of PIP2s with high water perme-ability (Section 5.2) in the endodermis could increase the radial hydraulicconductance and confer an efficient mechanism of regulating Lpr.

Additional support for PIP2s facilitating radial water transport acrossmembranes where the apoplast is blocked comes from aeroponicallygrown maize roots. Roots grown under these conditions develop a suber-ized exodermis and endodermis, in comparison with roots grown hydro-ponically (Hose et al., 2000; Zimmermann et al., 2000). Labeling ofZmPIP2;1 and ZmPIP2;5 by immunocytochemical localization was stron-gest in the epidermis, exodermis, and cortex of primary root tissue, corre-lating with the regions that were highly suberized (Hachez et al., 2006).Moreover, localization of ZmPIP2;5 in the epidermis exhibited polarity,with a stronger signal on the plasma membrane facing the external medium.The development of an exodermis in aeroponically grown maize rootstends to reduce Lpr compared with the less suberized hydroponic roots,


Table

2AQPtranscript(shownin

italics)andprotein

isoform

shighly

and/orpreferentiallyexpressedin

roots

Species

Isoform

Cell/tissue-specificlocalization

References

Arabidopsisthaliana

PIP2;2

Cortex,endodermis(highly

expressed),outerlayersofstele.

Javotetal.,2003

a PIP1;1,PIP1;2,PIP1;5

protein,

a PIP2;1,a PIP2;2,a PIP2;7,

TIP1;1,TIP1;2,TIP2;1

Expressionlevelsvariedam

ong

thedifferentstudies,reflecting

differentgrowth

conditions

(soilversushydroponic)and

growth

stage.

Alexanderssonetal.,2005;

Boursiacetal.,2005;

Danielsetal.,1996;Jang

etal.,2004;Santonietal.,

2003

SIP2;1

Primarilyexpressed

inroots,but

inlow

amounts.

JohansonandGustavsson,

2002

Craterostigmaplantagineum

CpPIPa(PIP1),CpPIPb(PIP1)

Mariauxetal.,1998

Hordeumvulgare

HvPIP2;1

Katsuharaetal.,2002

HvPIP2;1

protein

Allcellsnearthetip,but

epidermisandvascularbundles

inmaturingregion.

Katsuharaetal.,2003a

Juglansregia

JrPIP2;2

Sakretal.,2003

Lotusjaponicus

LIM

P2(N

IPprotein)

LIM

P1(TIP

protein)

Symbiosomemem

brane.

Rootsandsymbiosome

mem

brane.

Guenther

andRoberts,2000

PIP1-likeprotein

Henzler

etal.,1999

Lycopersiconesculentum

LeA

QP2

Werner

etal.,2001

Mesem

bryanthem

um

crystallinum

MIP-A

(PIP

protein)

Epidermisandendodermisof

youngroots,vasculature

of

mature

roots.Pericycleand

cortex,notapex.

Kirch

etal.,2000;Yam

adaand

Bohnert,2000

MIP-B

(PIP

protein)

Endodermis.

MIP-C

(PIP

protein)

Allcellsofelongationzone.

MIP-F

(TIP

protein)

Allrootcells.

(continued)

Table

2(continued)

Species

Isoform

Cell/tissue-specificlocalization

References

Nicotianaglauca

NgM

IP1(TIP),NgM

IP5(PIP)

Smartetal.,2001

Nicotianatabacum

aNtAQP1(PIP1b)

Transcripthighlyabundantin

roots.Roottipandmeristem.

Associated

withxylem.Protein

detectedin

exodermis,

endodermis,cortex,near

vascularbundles,xylem

parenchyma,andapicalregion.

Bielaetal.,1999;Ottoand

Kaldenhoff,2000;Siefritz

etal.,2001

Oleaeuropaea

OePIP1;1,OePIP2;1

Highestin

roots.

Secchietal.,2007

Orysasativa

rwc1

(PIP1)

OsPIP2;3,OsPIP2;4,

andOsPIP2;5

Lsi1(N

IP)

Predominantlyexpressed

inroots.

Silicontransporter

primarily

expressed

inroots,localizedat

theplasm

amem

braneofthe

distalsideoftheexo-and

endodermis.

Lietal.,2000

Sakuraietal.,2005

Maetal.,2006

Pisumsativum

PsPIP1;1,PsPIP2;1,PsTIP1;1

Highestin

roots.

Schuurm

ansetal.,2003

Populustrem

ulax

trem

uloides

PttPIP1;1,PttPIP2;2,PttPIP2;3,

PttPIP2;5

Preferentiallyexpressed

inroots.

Marjanovicetal.,2005

Raphanussativus

ag-VM23(TIP)

a PAQ1s(PIP1s)and

a PAQ2s

(PIP2s)

Growingrootsandtaproot.

Highestin

youngroots,butalso

highin

mature

taproots.

Sugaetal.,2001

RsPIP1andRsPIP2proteins

RsTIP

Vasculature

oftaproots,but

endodermisofseedling.

Alltissueoftaproots,epidermisof

seedlingroots.

Sugaetal.,2003

RsPIP2;1

Sugaetal.,2002

Spinaciaoleracea

pm28a(PIP2;1)

a SoPIP1;2

Highestin

roots.

Fraysseetal.,2005

Johanssonetal.,1996

154

VitishybridRichter110

PIP1;1

andTIP1

PIP2;1

andPIP2;2

Highestin

roottipandlateral

roots.

Highestin

roots.

Baiges

etal.,2001

Zea

mays

ZmPIP1;2,ZmPIP2;4

Highestin

mature

compared

to

elongatingtissue.

Hukin

etal.,2002

aZmPIP1;5

Preferentiallyexpressed

inroots.

Steleandcortex.

Gaspar

etal.,2003

ZmTIP1

Meristemsofprimaryandlateral

roots.Epidermisand

endodermis,also

xylem

parenchymaofelongation

zone.In

mature

rootshighest

inxylem

parenchyma.

Barrieu

etal.,1998;Chaumont

etal.,1998

ZmPIP1a,ZmPIP2a

ZmPIP2a—

rootspecific.

Chaumontetal.,2000

ZmPIP1;2

ZmPIP2;5

Xylem

parenchyma.

Cortex

andxylem

parenchyma.

Fetteretal.,2004

aZmPIP1;1,

a ZmPIP2;1,aZmPIP2;5,

a ZmPIP1;5

Developmentallyregulatedin

aeroponicallygrownroots,

greatestexpressionin

mature

primaryroottissue.ZmPIP2;1

andZmPIP2;5

labelingin

mature

zones

greatestin

epidermis,exodermis,and

cortex.Alsoin

opericlinal

plasm

amem

braneofepidermis

exposedto

externalmedium.

Hachez

etal.,2006

ZmPIP1;5

ZmPIP2;4,ZmTIP2;1–2;3

Primarilyexpressed

inroots.

Chaumontetal.,2001

ZmTIP2;3

Rootspecific.

Lopez

etal.,2004

aIndicates

expressionofboth

transcriptandprotein.

155

but does not appear to influence Lpc (Hose et al., 2000; Zimmermann et al.,2000). Therefore, it would have been interesting if Hachez et al. (2006) hadexamined whether the expression of these PIP2s only occurred in rootswith an exodermis.

Certain PIP1 homologues are also highly expressed in roots, but there isless correlation with tissue-specific expression (Table 2). PIP1;1 homologuehas highest expression associated with the apical meristem in Vitis R-110roots, but is associatedwith vascular tissue inmature radish roots and a varietyof cell types in tobacco roots (Baiges et al., 2001; Otto and Kaldenhoff, 2000;Suga et al., 2003). These differences may reflect different functional roles forPIP1s that vary with developmental stage, but because most PIP1s have lowwater permeability when expressed in Xenopus oocytes (Section 5.2), theymay serve some regulatory role through interaction with other PIPs (Fetteret al., 2004), or as osmotic or turgor sensors (Hill et al., 2004).

Some TIP homologues are specifically expressed in roots and no otherorgans (e.g., ZmTIP2;3, Lopez et al., 2004). Conversely, other TIPs areubiquitous, being expressed in all root cells (e.g., MIP-F, Kirch et al., 2000),suggesting their importance in ‘‘housekeeping’’ processes such as the osmo-regulation of the cytosol. Some studies have shown that the tonoplast has amuch higher Pf than the plasma membrane, which would allow the vacuoleto protect against deleterious cytoplasmic volume changes (reviewed byChrispeels et al., 2001; Maurel, 1997; Tyerman et al., 1999, 2002). This rolein osmotic equilibration is further supported by the observation that severalTIPs are predominantly expressed in tissue, such as the epidermis, thatpotentially experience the greatest fluctuations in apoplastic water potential(e.g., ZmTIP1, Barrieu et al., 1998, Table 2). However, according to Hillet al. (2004) the greater AQP activity in the tonoplast serves some functionother than buffering against rapid changes in cytoplasmic volume becausepressure changes induced through transpiration or osmotic perturbations inthe soil are not instantaneously transmitted through the apoplast. SeveralTIP1 homologues are also primarily expressed in root tips of grapevines(V. vinifera), radish, and maize (Table 2), implying a putative role in turgormaintenance that is required to drive the cell expansion process (Chaumontet al., 1998; Dolan and Davies, 2004).

5.3.2. Changes in AQP abundanceThe stages of protein synthesis provide mechanisms for controlling theamount of functional AQPs present in a membrane. The majority of studiesinvestigating changes in AQP abundance have focused on expressionlevels of AQP mRNA transcripts, but this does not always correlatewith abundance of the respective proteins. AQP gene expression or proteinabundance in roots changes both temporally and in response to environmen-tal stimuli, and the expression patterns are multifaceted, varying between


genes and treatment (Table 3). Some AQPs are upregulated by abioticfactors, whereas others are either downregulated or not affected.

One of the most comprehensive studies undertaken was conducted byJang et al. (2004), who analyzed the transcript levels of all PIPs expressed inroots and aerial parts of A. thaliana, in response to drought, salinity, cold, orABA treatments. No consistent expression patterns existed in roots. Droughtand low temperature downregulated a large number of PIPs in roots,including most PIP2 transcripts, whereas salinity had a stimulatory effect.In comparison, Boursiac et al. (2005) found that salinity generally depressedAQP gene expression, although the effect depended on the particulartranscript and the time of exposure to the salinity treatment. An increase inAQPs may facilitate water uptake under harsh conditions, without necessi-tating large osmotic or hydraulic gradients between soil, xylem, and shoot.Alternatively, if water uptake is low, the presence of AQPs may increase thebackflow of water into the dry soil and cause tissues to dehydrate. Therefore,it is critical where AQPs or their activities are localized.

In comparison with the downregulation of a large number ofAQP transcripts in response to chilling in A. thaliana and rice roots(Table 3), PIP1 and PIP2 transcripts, as well as PIP2 protein abundanceappear to increasewith cold acclimation ofwheat crowns, which is associatedwith improved freezing tolerance (Herman et al., 2006). An increase inPIP2 abundance possibly enhances water export to the apoplast to reduceintracellular injury from the development of ice crystals, although nophysiological measurements were undertaken.

The phytohormones ABA and gibberelic acid (GA) appear to beinvolved in facilitating transcription of PIPs and TIPs from different plantspecies (Table 3). ABA is involved not only in signaling and stomatal closureduring drought, but ABA also potentially enhances Lpr (Section 4).

Most PIP genes appear to be diurnally regulated (Table 3). PIP transcriptexpression gradually increases, peaking 2–8 h into the photoperiod(depending on the plant species) and then declines to a basal level at thestart of or during the dark period (Gaspar et al., 2003; Henzler et al., 1999;Lopez et al., 2003; Sakurai et al., 2005). However, peak transcript andprotein expression occur at midnight in barley (Hordeum vulgare) roots(Katsuhara et al., 2003a). Lopez et al. (2003) observed that the diurnalpattern continues during continuous darkness suggesting circadian regula-tion, although the amplitude of expression diminishes with time.

The changes in AQP expression during the day/night cycle couldcontribute to diurnally fluctuating Lpr (Section 4). Henzler et al. (1999)found that the diurnal fluctuation of AQP transcripts of L. japonicus rootscoincided with fluctuations in Lpr. The oscillation in transcript abundanceand Lpr was slightly offset, with transcript expression increasing in anticipa-tion of the light period, before changes in Lpr. Paradoxically, the perme-ability of root cell membranes did not vary diurnally. However, regulation


Table

3AQPtranscriptexpression(shownin

italics)and/orprotein

abundance

shownto

vary

withtime,development,or

environmentalstim

uli

Stim

ulus

Plant

species

Upregulated

Dow

nregulated

Referen

ces

Drought

Arabidopsisthaliana

PIP1;3,PIP1;4,PIP2;1,

PIP2;5


PIP2;3

PIP2;4,PIP2;6,

PIP2;7

Jangetal.,2004

Craterostigma

plantagineum

CpPIPa2,CpPIPa6,

CpPIPa7,CpPIPc

Mariauxetal.,1998

Glycinemax

GmPIP1,GmPIP2(effect

greater

formycorrhizal

plants)

Porceletal.,2006

Helianthusannuus

SunTIP7,SunTIP20

(transient)

SunTIP18

Sardaetal.,1999

Latucasativa

LsPIP1,LsPIP2

Porceletal.,2006

Mesem

bryanthem

um

crystallinum

McT

IP1;2

protein

(mannitolorsorbitol

induced)

Vera-Estrellaetal.,2004

Nicotianaglauca

NgM

IP2,NgM

IP3(TIPs),

NgM

IP4(PIP)

Smartetal.,2001

Oryzasativa

RWC1(PIP1)(m

annitol)

Lietal.,2000

Oryzasativa

OsPIP1;1,OsPIP1;2

(24hPEG

treatm

ent)

OsPIP1;3,allOsPIP2s

(24hPEG

treatm

ent)

Liu

etal.,2006

Phaseolusvulgaris

PvPIP1;1,PvPIP1;2,

PvPIP2;1

PIP2proteins

Aroca

etal.,2006

Raphanussativus

RsPIP2;1

protein

Sugaetal.,2002

Salinity

Arabidopsisthaliana


PIP1;4,PIP2s

PIP1;5,

Jangetal.,2004

Arabidopsisthaliana

PIP1;1

(24h60–100mM

NaC

l)

Martinez-B

allestaetal.,

2003

158

Hordeumvulgare

HvPIP2;1,HvPIP2;1

Katsuharaetal.,2002

Mesem

bryanthem

um

crystallinum

MIPC

(PIP2protein)

McT

IP1;2,(M

IPFprotein)

Kirch

etal.,2000;Vera-

Estrellaetal.,2004

Oryzasativa

RWC1(PIP1)

Lietal.,2000

Raphanussativus

RsPIP2;1

protein

Sugaetal.,2002

Nutrientstatus

Arabidopsisthaliana

AtTIP1;2,AtTIP2;1

and

AtTIP4;1

(N-

starvationinduced)

Liu

etal.,2003

Lotusjaponicus

PIP1-like(N

O3�

starvation)

Clarksonetal.,2000

Oryzasativa

Lsi1(continuoussilicon

supply)

Maetal.,2006

Oryzasativa

OsPIP1;1–1;3,

OsPIP2;2,OsPIP2;3,

OsPIP2;5,OsPIP2;7

(24hKþstarvation)

Liu

etal.,2006

Zea

mays

ZmPIP1;5

(NO

3�

inducedafterN

starvation)

Gaspar

etal.,2003

Cold

Arabidopsisthaliana

PIP2;5,PIP2;6


PIP2;2,PIP2;3,

PIP2;4;PIP2;7

Jangetal.,2004

Oryzasativa

OsPIP1;3

RWC1(PIP1),OsPIP1;1,

OsPIP1;2,

OsPIP2;1–2;6,

OsTIP1;1,OsTIP2;2

Lietal.,2000;Sakurai

etal.,2006

Triticumaestivum

PIP2b,PIP2protein

(coldacclim

ation)

Herman

etal.,2006 (continued)

159

Table

3(continued)

Stim

ulus

Plant

species

Upregulated

Dow

nregulated

References

Zea

mays

PIP1proteins

Aroca

etal.2005

Pathogen

Lycopersicon

esculentum

LeA

qp2(infectionby

parasiteCuscuta)

Werner

etal.,2001

Nicotianatabacum

TobRB7(root-knot

nem

atode)

Opperman

etal.,1994

Symbiosis

Glycinemax

NOD26protein

(N2fixingnodule)

Guenther

etal.,2003

Medicagotruncatula

MtAQP1(g-T

IP)

(mycorrhizal)

Krajinskietal.,2000

Populustrem

ulax

trem

uloides

PttPIP1;1,PttPIP2;3,

PttPIP2;5

(mycorrhizal)

Marjanovicetal.,2005

Phytohorm

ones

Arabidopsisthaliana

(ABAorGA)


PIP1;4,PIP2;2,

PIP2;3,PIP2;4

PIP1;5,PIP2;6

Jangetal.,2004;

Kaldenhoffetal.,1996

Lycopersicon

esculentum(IAA)

LeA

qp2

Werner

etal.,2001

Mesem

bryanthem

um

crystallinum

(ABA)

McTIP1;2

Vera-Estrellaetal.,2004

Nicotianatabacum

(ABAorGA)

NtAQP1

Siefritzetal.,2001

Phaseolusvulgaris

(ABA)

PIP1proteins

Aroca

etal.,2006

Developmental

Arabidopsisthaliana

AtTIP1;2,AtTIP2;1,

AtTIP4;1

Liu

etal.,2003

160

Mesem

bryanthem

um

crystallinum

MIP-A

toMIP-F

proteins

Kirch

etal.,2000

Nicotianatabacum

NtAQP1

OttoandKaldenhoff

2000;Siefritzetal.,

2001

Raphanussativus

RsPIP1,RsPIP2,PAQ2

(RsPIP2protein)

Sugaetal.,2001

Diurnalcontrol

Hordeumvulgare

HvPIP2;1,HvPIP2;1

protein

Katsuharaetal.,2003a

Lotusjaponicus

PIP1a

Henzler

etal.,1999

Oryzasativa

OsPIP1;2,OsPIP1;3,

OsPIP2;3,OsPIP2;4,

OsPIP2;5,OsTIP1;2,

OsTIP2;1

Sakuraietal.,2006

Raphanussativus

PAQ1andPAQ2

proteins

Sugaetal.,2001

Zea

mays

ZmPIP1;1,ZmPIP1;5,

ZmPIP2;1,

ZmPIP2;5,ZmPIP1

protein,ZmPIP2

protein

Gasparetal.,2003;Lopez

etal.,2003

Light

Arabidopsisthaliana

PIP1b(w

hiteorblue

lightactivated)


TIP1;1,TIP2;1,

TIP2;2

Kaldenhoffetal.,1996;

Sato-N

araetal.,2004

Dark

Arabidopsisthaliana

TIP1;1,TIP1;2,

TIP2;2,PIP1;2,

PIP2;1,PIP2;3

Sato-N

araetal.,2004

Zea

mays

ZmPIP1protein

ZmPIP2protein

Lopez

etal.,2003

Circadian

Zea

mays

ZmPIPs

Lopez

etal.,2003

161

in Lpr could be controlled at the endodermis or stele, and the cell pressureprobe measurements were only conducted on cortical cells.

Lopez et al. (2003) observed a positive correlation between the amount ofZmPIP2 proteins (but not PIP1 proteins) and the rate of water flow throughde-topped maize seedlings. Under an osmotic gradient, stimulated waterflux from the root system, but continuous darkness stopped sap flow andcaused the relative amount of PIP2 proteins to decline. Other regulatorymechanisms also appeared to be involved, as the abundance of PIP2 proteinsbegan to increase before the start of the light period and before the start ofsap-flow. Katsuhara et al. (2003a) also correlated AQP protein abundancewith Lpr of barley roots. Contrary to ZmPIP2 proteins, HvPIP2;1 proteinaccumulated during late evening and peaked at night, following the samediurnal oscillation as Lpr. HvPIP2;1 was detected in most cells 2 mm fromthe tip, but was localized to the epidermis, outer cortical layer and sur-rounding the vascular cylinder, 50 mm from tip. Katsuhara et al. (2003a)speculated that posttranslational modifications of HvPIP2;1 could also bemodulating Lpr because the amplitude of the oscillation in Lpr was greaterthan that of HvPIP2;1 protein abundance. The studies of Katsuhara et al.(2003a) and Lopez et al. (2003) are also important because they demonstratethat the turnover rate of AQPs is rapid. For example, the abundance ofZmPIP1 proteins in maize roots can increase more than 20-fold within 4 h(Lopez et al., 2003).

Gaspar et al. (2003) also found that ZmPIP1;5 transcript varied diurnally,with high expression in the stele and cortex during the day and lowerexpression during the night. However, analogous to HvPIP2;1, the expres-sion of ZmPIP1;5 during the night was limited to the epidermis. The cellwalls of the epidermis, or (if present) the exodermis, of some plant speciesare suberized so expression of AQPs in the epidermis may facilitate wateruptake in the symplast when the apoplast is blocked. However, this does notexplain why the localized expression of ZmPIP1;5 changes diurnally. Ifepidermal expression during the evening is a common feature of certain PIPisoforms, it raises the question about possible involvement in the redistribu-tion of soil water by plant roots. The roots of several tree species ‘‘redistrib-ute’’ soil water during the night by absorbing water from the wetter parts ofthe soil profile and releasing it to the dryer regions (Burgess et al., 1998).The water is then taken up during the day, when transpiration resumes. Theexpression of AQPs in the epidermis of roots, during the night, couldmediate this hydraulic redistribution by either enhancing the backflow ofwater into the dry soil and/or enhancing the uptake of water from wet soil.

5.3.3. Posttranslational regulationWhen extrapolating information on transcription levels, care is needed asprotein abundance or activity is not necessarily altered (reviewed byChrispeels et al., 2001; Tyerman et al., 2002). Translation of the mRNA


transcript into protein may be delayed or inhibited until a signal is received,or the RNAmay require molecular changes after transcription. The amountof protein translated often does not correlate with the amount of transcriptexpressed (Alexandersson et al., 2005; Aroca et al., 2005; Kirch et al., 2000;Lopez et al., 2003; Suga et al., 2001). Once the message in mRNA istranslated, a polypeptide chain forms, but the protein may still not functionas a water channel. The majority of AQPs have putative glycosylation andphosphorylation sites where attachment of carbohydrate and phosphategroups, respectively, may be required to enable correct folding of theprotein and/or insertion into the membrane. Little is known about endo-cytosis processes in plants, but hormones and environmental conditionscan regulate the internalization and recycling of plasma membrane pro-teins, including some AQPs (Murphy et al., 2005). Santoni et al. (2000),Chrispeels et al. (2001), and Tyerman et al. (2002) have reviewed traffickingof AQPs to and from the membrane, as a regulatory mechanism of activity.Since then, Vera-Estrella et al. (2004) demonstrated in Mesembryanthemumcrystallinum that McTIP1;2 becomes glycosylated, in response to osmoticperturbation, and is trafficked to other compartments, where it thenbecomes de-glycosylated. Activation of the cAMP-signaling pathway andphosphorylation of the protein by protein kinase are also involved inredistribution of this AQP. An innovative proteomic study by Santoniet al. (2003) demonstrated that individual PIP isoforms can be present indifferent forms and at least one of these forms was due to posttranslationalmodification by phosphorylation.

Methylation is also a potential posttranslational regulatory mechanism ofAQP function (Santoni et al., 2006). Several residues of the N-terminal tailof PIPs can be methylated, which may influence protein stability andsubcellular localization. However, expression of PIP2;1 with altered meth-ylation sites in A. thaliana suspension cells did not significantly change theirosmotic water permeability (Santoni et al., 2006).

5.3.3.1. AQP gating Once the AQP is located in the membrane, openingand closing of the channel can then control water flow through the pore.Phosphorylation, cytoplasmic pH and heavy metals directly control AQPgating, either through conformational changes in the shape of the pore ordirect blockage of the pore. Ca2þ also potentially acts by direct or indirectobstruction of the pore (Alleva et al., 2006; Gerbeau et al., 2002). Othermechanisms that affect water permeability, speculated to be via interactionwith the AQP pore, include mechanical stimuli and osmotic pressure (Wanet al., 2004; Ye et al., 2005).

5.3.3.1.1. Phosphorylation By site-directed mutagenesis, Johansson et al.(1996, 1998) demonstrated that the phosphorylation/dephosphorylationof two serine residues regulates PM28A (now called SoPIP2;1) of spinach


(Spinacia oleracea) leaf, which increases/decreases Pf, respectively. In vivo,phosphorylation of Ser-274 depends on the water potential of the leafapoplast and based on their observations, Johansson et al. (1998) developeda model for the regulation of flow through PM28A. During ambientconditions, the leaf apoplast has a high water potential and cell turgor ishigh. Submicromolar Ca2þ activates a membrane-associated protein kinasethat phosphorylates PM28A, on Ser-274, and the water channel opens.When the leaf experiences water deficiency, cell turgor begins to fall, theprotein kinase inactivates (possibly by closure of a stretch-activated Ca2þchannel) and dephosphorylation of PM28A closes the pore. It is likely thatwater-potential-induced phosphorylation regulates water channel activityin roots as well as leaves, as PM28A is predominantly expressed in roots andSer-274 is conserved in all PIP2s.

Using X-ray crystallography Tornroth-Horsefield et al. (2006) made aremarkable breakthrough by identifying the phosphorylation-inducedstructural mechanism that gates SoPIP2;1 in spinach. Plant PIPs typicallyhave a longer Loop D (four to seven amino acid residues) that folds underthe protein and occludes the pore. Phosphorylation of Ser-115 and Ser-274causes the ‘‘cap’’ of LoopD to be displaced so that the pore is open. Throughmolecular modeling, Tornroth-Horsefield et al. (2006) also demonstratedhow Ca2þ and protonation of His 193 may regulate the gating of PIPs(Section 5.3.3.1.2).

The increasing evidence implies that phosphorylation of AQPs is animportant regulator of water channel activity. The water channel of radishRsPIP2;2 and soybean NOD26 is activated by phosphorylation (Guentheret al., 2003; Suga and Maeshima, 2004). Phosphorylation of RsPIP2;2 couldbe a fundamental factor influencing Lpr, particularly during changes inexternal osmotic pressure, since this AQP is highly expressed in roots(Table 2). Phosphorylation of NOD26 is dependent on the developmentalstate of the nodule, with maximum activity in mature nodules (Guentheret al., 2003). Contrary to their effects on PM28A, drought (and salinity)enhances the phosphorylation of NOD26 in planta, although a concurrentincrease in water permeability was not assessed in planta, phosphorylationinduced by okadaic acid increased Pf in Xenopus oocytes (Guenther et al.,2003). However, decreasing the external osmotic potential reduced Pf ofmembranes vesicles isolated from the peribacteriod membranes (PBM)of soybean root nodules, in which NOD26 is highly abundant (Vandeleuret al., 2005). The response of NOD26 in PBM vesicles is consistent withJohansson’s model ( Johansson et al., 1998), but conflicts with the findings ofGuenther et al. (2003) and highlights the importance of measurements onnative membranes.

Temmei et al. (2005) demonstrated that phosphorylation of PIP1 pro-teins was involved in the cooperative regulation of water channel activitywith PIP2 proteins. While phosphorylation did not affect the formation of


heterotetramers, phosphorylation of Ser-131 of MpPIP1;1 (in Mimosapudica) activated the water channel activity of MpPIP2;1.

Presently, no studies have investigated phosphorylation-regulated Lpr inresponse to abiotic perturbations in root systems. However, Kamaluddinand Zwiazek (2002b) applied exogenous ethylene to hypoxic or aeratedP. tremuloides seedlings, which increased Lpr. They interpreted this responseas an increase in water channel activity by phosphorylation, as ethyleneinduces protein phosphorylation with involvement of Ca2þ-dependentprotein kinases. In plants that are waterlogged, hypoxia strongly stimulatesthe biosynthesis of ethylene, but at the same time tends to reduce Lpr (Drew,1997; and Section 7.3), which conflicts with Kamaluddin and Zwiazek’sideas. Experiments inhibiting ethylene biosynthesis or action, or phosphor-ylation during hypoxia, or stimulating phosphorylation in the absence ofethylene during hypoxia, may resolve this issue.

5.3.3.1.2. CytoplasmicpH Studies on the vacuoles and plasmamembranesof Beta vulgaris, and the plasma membranes of A. thaliana, have shown thatcytoplasmic pH affects the gating of AQPs and that acidification of the cytosolreduces the membrane permeability to water (Alleva et al., 2006; Amodeoet al., 2002; Gerbeau et al., 2002; Sutka et al., 2005). This is important asrespiratory inhibitors, anoxia, and some other abiotic perturbations induceacidification of the cytosol (Dat et al., 2004; Felle, 1987, 2005).

The connection between anoxia-induced cytoplasmic acidification andAQP gating by internal pH is particularly relevant in understanding thewaterlogging-induced reductions in Lpr (Section 7.3). A pioneering studyby Tournaire-Roux et al. (2003) identified a histidine residue conserved inall PIPs, which is a major site for cytosolic pH sensing. Tournaire-Rouxet al. (2003) also found that a group of charged amino acid residues on LoopD of the protein manipulate AQP gating by controlling either the confor-mation and/or the conductance of the pore, in response to cytosolic pro-tons. When a cell experiences anoxia, the cytoplasm acidifies due to anaccumulation of by-products from anaerobic respiration. The histidine ofPIPs becomes protonated, which closes the water channel and so reducesthe permeability of the membrane to water (Tournaire-Roux et al., 2003).

It should also be noted that during O2 deficiency root cells leak ions,which is assumed to be due to depolarization of the plasma membrane thatstimulates the opening of ion channels (e.g., wheat; Greenway et al., 1992;Zhang and Tyerman, 1997). Given the acid-induced gating of the mam-malian AQP6 (Hazama et al., 2002), anoxia could stimulate similar ionpermeability in analogous AQPs in plants.

5.3.3.1.3. HeavyMetals Mercuric chloride has been widely used to dem-onstrate the function of AQPs in isolated native membranes and in planta.Hg2þ binds to cysteine residues on the protein and acts either by directobstruction of the pore or by conformational change (Section 5.1, Barone


et al., 1997, 1998). Blocking water channels reduces the membrane perme-ability to water and this has been demonstrated in Xenopus oocyte swellingassays, membrane vesicles, isolated protoplasts, and intact cells (Daniels et al.,1996; Kaldenhoff et al., 1998; Niemietz and Tyerman, 1997; Zhang andTyerman, 1999).

AQP inhibitors are valuable because they provide an estimate of AQPactivity in real time, without destruction of the tissue and can be used toexamine AQP activity in relation to changing environmental conditions.Mercury is nonspecific (cannot target individual isoforms or may affectother channel proteins), may be phytotoxic and may reduce cellular metab-olism and other biochemical processes (reviewed by Santoni et al., 2000;Tyerman et al., 1999), but despite all these caveats a reduction in cell waterpermeability indicates a decrease in AQP activity.

Niemietz and Tyerman (2002) demonstrated that gold and, in particular,silver ions inhibit AQPs more effectively than mercury. Silver ions reducedthe water permeability of membrane vesicles from different organisms, toalmost the predicted inherent water permeability of the lipid bilayer. Theeffect of silver was not reversible and the mechanism of inhibition isunknown, but Niemietz and Tyerman (2002) reasoned that Agþ andAu3þ react with sulfhydryl groups of cysteine residues, analogous to mer-cury. Silver is considered less toxic than mercury and provides a promisingalternative for AQP inhibition studies (Niemietz and Tyerman, 2002).Confirming Niemietz and Tyerman’s observations, gold and silver inhibitedthe activity of AQP1 in erythrocytes and epithelial cells from mice, but to asimilar extent as mercury (Yang et al., 2006).

Despite Tazawa and Iwasaki (1996) demonstrating that Zn2þ reducedLpcof Chara corallina cells, Zn2þ and other transition elements did not signifi-cantly reduce the water permeability of membrane vesicles (Niemietz andTyerman, 2002). However, Tazawa and Iwasaki (1996) used a high concen-tration of Zn2þ (5 mM) and the speed of the inhibition on Lp depended onthe pH of the external medium, suggesting that AQP activity may have beenindirectly affected.

5.3.3.1.4. OtherAQPgatingmechanisms Meinild et al. (1998) found thatAQP pore dimensions are not fixed and that lowering the temperaturecaused the pore to narrow, which increased the reflection coefficient forvarious solutes. Wan et al. (2004) speculated that mechanical stimulus closesAQPs, either by creating tension that collapses the pore or causes a con-formation change in the pore shape. However, Ca2þentering throughstretch-activated ion channels may also indirectly close AQPs during pres-sure pulses. Large positive or negative pressure pulses (>0.1 MPa) inducedby the cell pressure probe reduced the water permeability of cortical cellsin maize roots (Wan et al., 2004). The reduction was transient with 0.1–0.2MPa pressure pulses, but pulses >0.2 MPa were only reversible by theaddition of high concentrations of ABA. Wan et al. (2004) verified that the


reduction in water permeability was by a reduction in AQP activity, as thosecells whose water permeability had been reduced by large pressure pulseswere unaffected by mercury. This finding by Wan et al. (2004) is importantas the cell pressure probe is widely used for measuring the water permeabilityof cells in intact tissue. Future measurements with the cell pressure probeshould be restrained to applying small pressure changes (pressure relaxations)to prevent confounding effects on cell water permeability.

Osmotic pressure has also been suggested to gate AQPs through dehydra-tion of the pore (Ye et al., 2004, 2005). The observation that the permeabilityof the membrane to water decreases with increasing external solute concen-tration has been reviewed byTyerman et al. (2002) andVandeleur et al. (2005).They speculate that high osmotic pressure could not only restrict the porethrough dehydration of the protein but also could create tension in the centralvestibule of the tetramer causing conformation changes of themonomers. Theinhibition of AQP activity by high external solute concentration has relevanceto water flow through root systems, as salinity tends to reduce Lpr (Section 4).Salinity appears to stimulate the expression of AQPs in A. thaliana roots ( Janget al., 2004), but this will probably be ineffectual at influencing Lpr if highsolute concentrations close these AQPs.

6. The Role of AQPs in Root Water Transport

6.1. Inhibition studies

The inhibition of water flow by mercuric chloride has illustrated thatAQPs are an important component in regulating water transport across theroots of some species (Barrowclough et al., 2000; Maggio and Joly, 1995;Martinez-Ballesta et al., 2000; Martre et al., 2001; Quintero et al., 1999; Wanand Zwiazek, 1999). The use of mercuric chloride has revealed that AQPs canaccount for up to 90% of total root water flow either through occlusion of thewater channel or indirect inhibition by lowered metabolism (Martre et al.,2001 and references therein).

The employment of mercury, in combination with an abiotic perturba-tion, has confirmed the inhibition of AQP activity. For example, salinityreducedLpr (based on flux per gram rootweight) ofCucumis melo andCapsicumannuum roots by 80% and 100%, respectively (Carvajal et al., 1999, 2000).Mercuric chloride reducedLpr of control plants, to a similar extent as the effectof salinity, but barely influenced Lpr of salinity-treated roots (Carvajal et al.,1999, 2000). Similarly, nutrient deficiency reduced Lpr of wheat roots by 70–80%,which was not further inhibited bymercury, but mercury reduced Lpr ofnutrient-sufficient roots by 63% (Carvajal et al., 1996).

Measurements of the radial hydraulic conductivity after removal of tissuelayers and application of mercury demonstrated the varying activity of AQPsin different regions of roots of cacti and desert succulents (Martre et al., 2001;


North et al., 2004). Lpr decreased with soil drying in the distal and mid-rootregions of Opuntia acanthocarpa, with no additional reduction by mercury(Martre et al., 2001). In wet soil, mercury reduced Lpr of the distal region, butnot the mid-root region. However, mercury may not have fully permeatedthe mid-root region. After removing the cortex and periderm, mercuryreduced the radial hydraulic conductivity of the stele, which accounted foralmost 75% of Lpr (Martre et al., 2001). In an analogous study onAgave desertiroots, AQPs were found to be active external to the stele, being important inthe epidermis/exodermis of the basal region, but more important in theendodermis of the distal region (North et al., 2004).

Wan and Zwiazek (1999) and Siemens and Zwiazek (2003) used anapoplastic permeant dye to demonstrate that mercury inhibits the water fluxfrom P. tremuloides roots, but the relative contribution of the apoplasticpathway increased. The relative contribution of the apoplastic pathway alsoincreased when AQP activity was inhibited by mercury treatment inA. thaliana roots (Martinez-Ballesta et al., 2003). The contribution of thedifferent pathways to Lpr could also be estimated using the cell pressureprobe to measure the effects of mercury on the water permeability of rootcells and comparing them with measurements of root hydraulic conductiv-ity. However, such comparisons in the literature are rare. Barrowcloughet al. (2000) combined cell pressure probe and root potometer measure-ments, to identify the dominance of apoplastic transport in the young regionof onion roots, but AQPs facilitated water flow in the mature root regionscontaining a suberized exodermis. Although Zhang and Tyerman (1999)did not measure Lpr, the inhibition of Lpc by mercury was similar to thatobserved by Carvajal et al. (1996), in whole roots, suggesting a significantcontribution of the cell-to-cell pathway to Lpr of wheat roots. The combi-nation of CPP, RPP, and mercury treatment demonstrated the contributionof the radial pathways to flow in wheat and lupin roots (Bramley, 2006).Mercury treatment reduced Lpc to 15–20% of untreated cells, indicatinga significant influence of AQPs in membrane water permeability. Thisinhibition at the cellular level was associated with a similar inhibition inLpr (40–50% of control roots) of wheat roots, indicating that a significantproportion of radial water flow occurs through the cell-to-cell pathway andAQPs may be involved in regulating that flow. However, despite thepresence of mercury sensitive AQPs in lupin roots, mercury did not inhibitLpr (Bramley, 2006) suggesting that radial water flow may be predominantlyapoplastic. Moreover, the study by Bramley (2006) implies that AQPs mayhave other, as yet undiscovered, functional roles in roots.

AQP activity in roots may be an important component in the refilling ofembolized vessels in shoots of grapevine. The inhibition of root water flowby mercuric chloride impaired the recovery of shoot hydraulic conductanceafter a water deficit (Lovisolo and Schubert, 2006). Despite using a highconcentration of mercury, Lovisolo and Schubert (2006) were able to


reverse the inhibition of root hydraulic conductance and consequently theshoots rehydrated. The mechanism of repair after drought is unknown, butembolized vessels in grapevines refill in spring under root pressure(Scholander et al., 1955; Sperry et al., 1987). Inhibition of AQP activity inroots may therefore have some regulatory effect on root pressure, althoughdue to the nonspecificity of mercury nutrient transport would also havedecreased, which would affect root pressure.

6.2. Expression and transformation studies

Molecular transformation techniques creating AQP-deficient or overex-pressing mutants have corroborated the mercury inhibitory experiments inassessing the function of AQPs in plants. Knocking out NtAQP1 fromtobacco reduced Lpr by 58% compared with control plants, without alteringplant morphology (Siefritz et al., 2002). Conversely, downregulation ofPIPs in antisense A. thaliana plants increased the root mass, but had noeffect on root water uptake (Kaldenhoff et al., 1998; Martre et al., 2002).Kaldenhoff et al. (1998) speculated that antisense plants compensated for thelack of PIPs by increasing the absorption area of the root system. In supportof this, root systems of transgenic tobacco plants overexpressing PIP1b wereable to support a larger shoot system (Aharon et al., 2003).

Knocking out NtAQP1 from tobacco or PIP1s from A. thaliana reducedthe tolerance of antisense plants to a water deficit (Kaldenhoff et al., 1998;Siefritz et al., 2002), which is surprising given that PIP1 genes seem to bedownregulated under adverse conditions (Table 3). However, transgenictobacco and rice plants over expressing PIP1s and HvPIP2;1, respectively,were also less tolerant of drought or salinity (Aharon et al., 2003; Katsuharaet al., 2003b). There appears to be a fine balance between the abundance ofAQPs, root morphology, and water transport, so that some plants compro-mise performance under favorable conditions to minimize deleteriouseffects in the event of adverse conditions.

Javot et al. (2003) knocked out PIP2;2 that is predominantly expressed inA. thaliana roots. Lpc of cortical cells was almost one-third less for mutantsthan wild-type plants, but this reduction did not translate into an equivalentreduction in Lpr. The Lpr of detopped roots was only reduced by 14% underan osmotic driving force (natural sap exudation) compared with zeroreduction under a hydrostatic driving force (pressurized root system).Javot et al. (2003) interpreted their observations according to the compositetransport model, that is under a hydrostatic driving force, water flow waspredominantly via the apoplast (Section 2.3.1). However, expression ofPIP2;2 is predominantly localized in the endodermis (Table 2), implyingan important function of this AQP in this tissue. Manipulating AQP activityin the endodermis would be a geometrically efficient method of controllingLpr, given the decreasing surface area for water flow toward the root


axis. If the endodermis is suberized, then the apoplastic pathway may beblocked and water would have to cross the plasma membrane at least twice(i.e., through AQPs) to reach the apoplast of the stele. Therefore, evenunder a hydrostatic driving force there should be some transcellular flow.It would have been useful, if Javot et al. (2003) had examined the anatomy ofthe roots, not only to confirm the above speculations, but more impor-tantly, to ensure that A. thaliana mutants did not compensate for an absenceof PIP2;2 by some other mechanism.

The analysis of AQP expression in conjunction with measurements ofthe water permeability of cells or protoplasts has been used to identify thecontribution of AQPs to membrane permeability. Hukin et al. (2002)observed a higher Lpc of cells in mature compared with elongating tissueof maize roots, which was correlated with a greater expression of PIP genes.They suggested that AQPs are required in mature tissue to mediate radialwater flow because there was also less symplastic connection between cellsvia plasmodesmata in mature tissue. Suga et al. (2003) measured Pf of cellswith different AQP protein content. The osmotic water permeability ofcortical or endodermal protoplasts, isolated from radish roots, was the samedespite the endodermis having much greater expression levels of PIP1,PIP2, and a TIP. High Pf of protoplasts isolated from young rape (Brassicanapus) and flax (Linum usitatissimum) roots were considered to be related tohigh amounts of PIP1 and PIP2 proteins, but this was not true for twowheat genotypes that had low Pf, despite the presence of PIPs (Morillon andLassalles, 2001). Interestingly, wheat species with low Pf values were moretolerant of severe water deficit and were still able to germinate, suggestingstronger regulation of AQP activity may mediate the effects of water deficit,at least in germinating seeds. It should also be noted that the Pf values forwheat protoplasts and plasma membranes vesicles are much lower than insitu values obtained with the CPP (Bramley, 2006; Niemietz and Tyerman,1997; Zhang and Tyerman, 1999), indicating that the isolation proceduresmay affect AQP activity. Gerbeau et al. (2002) demonstrated that plasmamembrane vesicles isolated in the presence of chelating agents had higher Pfthan those isolated in the presence of divalent cations.

6.3. The contribution of AQPs to radial water flow

Given the ubiquitous nature, diversity, and high expression levels of manyAQPs, it is tempting to speculate that they have an important role inwater flowthrough the plant. However, AQPs will only have an influence on Lpr if asignificant proportionofwater flowoccurs by the cell-to-cell pathway (Steudle,1997). In some species, water appears to flow predominantly via the apoplast,for example maize and lupins, despite an abundance of AQPs expressedin the roots (Bramley, 2006; Chaumont et al., 2001; Steudle et al., 1987).


Therefore, the function of AQPs in these plants is an enigma, although thecontributionof the radial pathways to flowmayvaryunder different conditions,asGibbs et al. (1998b) thought that the contributionof the apoplast inmaize hadbeen overestimated.

There is the potential for finer control of Lpr, mediated by AQP activity,when water flows via the cell-to-cell pathway. The apoplast only constitutesa small fraction of the root cross-sectional surface area, so the cell-to-cellpathway could potentially transport larger volumes of water. Additionally,AQPs create parallel pathways for water flow across a membrane and there-fore their respective conductances are additive, so their abundance can have adramatic effect on Lpc. Increasing Lpc requires a lower gradient to drive thesame amount of flow. Under conditions of high transpiration, increasing theabundance of AQPs could considerably increase Lpr and hence avert largetensions in the xylem. Unless negative pressure (tension), generated bytranspiration and transmitted throughout the apoplast, causes a conforma-tional change in the protein and closes the pore, water flow through the cell-to-cell pathway would be advantageous. Measurements demonstrating thatabiotic perturbation, respiratory inhibitors, and mercury reduce hydrostati-cally induced water flux, point toward a significant proportion of root waterflow occurring via the cell-to-cell pathway, in many species (Section 4).

7. Waterlogging

Waterlogging occurs when the infiltration of water from rainfall orflooding is greater than the rate of subsurface drainage and evapotranspira-tion, leading to saturation of the soil. The frequency and duration ofwaterlogging, and the depth of soil saturation, depend not only on the soilproperties, but also on location in the landscape and climate. For example,during winter in Western Australia, when rainfall exceeds pan evaporation,the soil profiles susceptible to waterlogging often saturate close to the soilsurface, for days to several weeks at a time (Cox and McFarlane, 1995;Tennant et al., 1992; Zhang et al., 2004). Waterlogging is a significantenvironmental constraint for crop growth and can ultimately reduce yieldsby as much as 80% (Dracup et al., 1992; McFarlane and Williamson, 2002).

7.1. Effect on O2 in the rhizosphere

Oxygen diffuses 10,000 times slower in water than in air (Grable, 1966).Consequently, when the soil is saturated the O2 concentration in the rhizo-sphere declines due to respiring roots and microorganisms (Armstronget al., 1991; Cannell and Jackson, 1981). Greater respiration rates causedby high temperatures lead to a greater consumption of O2 and a more rapiddecline in O2 concentration in waterlogged soils over time (Belford, 1981


and Drew, 1992). The lack of soil aeration leads to the formation of manybiochemically reduced compounds that can accumulate to concentrationsthat are phytotoxic, but a low O2 supply is the primary factor detrimental toplant growth (Drew, 1992).

Even in winter, when the average daily temperatures are low, therhizosphere can become hypoxic (low O2) within a few days of water-logging. Barrett-Lennard et al. (1987) used specially designed plots tocontrol waterlogging in the field, at a nonplanted site in Western Australia.Waterlogging gradually reduced the O2 concentration in the soil profile to�4% O2 (0.05 mol m�3) within 4–8 days of waterlogging, depending onthe depth in the profile. The decline in O2 would be more rapid in croppedfields, depending on root density and root depth (Drew, 1992).

7.2. Effect on root growth

The lack of O2 causes respiration to become anaerobic, which affectsmetabolism and leads to a cascade of biochemical and physiological changesthat may ultimately cause cell injury or death (Drew, 1992). Aerobicrespiration switches over to glycolysis, once the O2 supply drops belowthe critical oxygen pressure (COP). The COP varies depending on thespecies but at 25 �C is generally less than 10% O2 (Drew, 1997).

Glycolysis only produces one-eighteenth of the energy of aerobic respi-ration and therefore growth ultimately becomes inhibited (reviewed byDrew, 1992). In general, roots are more adversely affected than shoots(Davies et al., 2000a; Trought and Drew, 1980). Because of their highrates of O2 consumption the root apical meristems are the most sensitiveparts of the root to O2 deficiency (Drew, 1992). In maize roots the COP forthe root tip is equal to, or slightly greater than the concentration of O2 inequilibrium with air (Saglio et al., 1984). The COP of wheat roots is 0.16mol m�3 O2 in the apical region (0–2 mm from the tip) compared with aCOP of 0.05 mol m�3 O2 for mature tissue (10–12 mm from the tip)(Thomson et al., 1989). Peak water uptake in cereal roots occurs behindthe root tip (Section 3.2), so growth should be inhibited before wateruptake is affected during O2-deficient conditions.

The effects of waterlogging and/or O2 deficiency depends on the speciesand the duration of submergence, with the complete cessation of rootgrowth or even root death in the more sensitive species (Cannell andJackson, 1981; Grable, 1966). Waterlogging or hypoxia reduces the growthof wheat roots and the extent of the inhibition depends on the genotype(Huang and Johnson, 1995; Huang et al., 1994; Thomson et al., 1992).Seminal roots of wheat appear to be more adversely affected by water-logging than nodal roots, which often increase in abundance and continueto elongate (Huang et al., 1994; Thomson et al., 1992; Trought and Drew,1980; Wiengweera et al., 1997).


Based on their growth response, yellow lupin was more tolerant towaterlogging than other lupin species (Broue et al., 1976; Davies et al.,2000a,b). Narrow-leafed lupin is very sensitive to excess water and mild O2

deficiency. Watering to 130% field capacity only reduced the air-filledporosity from 28% to 21%, but depressed root growth (White andRobson, 1989). The air-filled porosity is the proportion of the bulk soilvolume filled with air and the critical value considered to affect plant growthis <10% (Grable, 1966). By reciprocal grafts of roots and shoots of narrow-leafed lupin with yellow lupin, Davies et al. (2000c) identified that it wasthe root system of lupins that conferred the genotypic tolerance towaterlogging.

Wheat is considered to be sensitive to waterlogging, but anecdotalevidence in the field indicated that wheat was more tolerant to waterloggingthan lupin, which was confirmed in a glasshouse study (Bramley, 2006;Dracup et al., 1992). Lupins have thick taproots (1- to 2-mm diameter),which would require a higher rate of O2 supply from the rhizosphere fornormal functioning, compared with thin wheat roots (Dracup et al., 1992).Wheat and lupin roots also differ in their waterlogging tolerance due toanatomical changes that occur during waterlogging. The tolerances ofdifferent wheat genotypes to hypoxia were associated with changes inroot system anatomy that increased the internal aeration of roots (Boruet al., 2003; Erdmann and Wiedenroth, 1986; Huang et al., 1994; Thomsonet al., 1992).

An increase in internal aeration or root porosity, in response to hypoxia,is a common feature of many wetland species including rice and somedryland species including the cereal crops Hordeum marinum, maizeand wheat, and the legume species soybean, cowpea (Vigna unguiculata) andLotus species (Colmer, 2003; Evans, 2003; Thomas et al., 2005 and referencestherein). Hypoxia stimulates the development of gas spaces, called lysigenousaerenchyma, through ethylene-induced cell death (reviewed by Evans,2003). In roots, aerenchyma usually forms in the cortex and these air spacesare connected longitudinally, creating a pathway for the diffusion of gases,from the aerial parts of the plant to the roots (Colmer, 2003; Evans, 2003).The effectiveness of internal aeration depends on root porosity (i.e., theamount of aerenchyma), respiration, and radial O2 loss (ROL) to the soil(Armstrong et al., 1991). Many wetland species, such as rice, develop abarrier in the epidermis or outer hypodermal layers that reduces the loss ofO2 from the root to the soil (Colmer, 2003). The nature of the barrier isunresolved but may involve tightly packed cells and wall modifications suchas thickening and lignification (Armstrong et al., 2000).

Waterlogging and/or lowO2 supply stimulates the development of aeren-chyma in wheat roots, predominantly in nodal roots, although the seminalroots of some genotypes also increase their root porosity (Erdmann et al., 1986;Huang and Johnson, 1995; Huang et al., 1994; Thomson et al., 1992;


Trought and Drew, 1980; Wiengweera et al., 1997). Wheat roots do notappear to develop a tight barrier to ROL (Colmer, 2003), so even withaerenchyma the internal aeration may be inefficient. Without a barrier toROL,O2 can diffuse across the cortex in the basal region of the root and be lostto the soil environment, so that the root tips have insufficient O2 to maintainviable meristems. Root tips have tightly packed cells with highmetabolic ratesand few air spaces. It is therefore not surprising that the root tips are the mostsensitive part of the root when the O2 supply becomes limited. Modeling thediffusion of O2 within graminaceous roots Armstrong et al. (1991) demon-strated that even with the development of aerenchyma, root growth wouldonly extend into anaerobic media to a depth of �30 cm. In addition, theypredicted that the stele would become anoxic before the cortex, which waslater experimentally confirmed in maize roots (Darwent et al., 2003; Gibbset al., 1998a).

The majority of research has focused on the effects of waterloggingduring the waterlogging event. Far less is known about recovery, whenthe wet soils have drained. Dracup et al. (1992) suggested that wheat rootsmight resume growth more rapidly than lupins, based on observations indifferent field studies. Yellow lupin roots are better at recovering afterwaterlogging than narrow-leafed lupin, but only when waterlogged at anearly stage of growth (Davies et al., 2000a). In a glasshouse study, Bramley(2006) found that wheat roots survived a waterlogging event better thanlupins and recovery in root growth was more rapid, with a significantamount of growth resuming from the apical region. In comparison,narrow-leafed lupin roots died with no subsequent recovery. Althoughthe majority of the root system of yellow lupin roots died, root growthresumed from the basal region, and this was associated with the weakdevelopment of aerenchyma (Bramley, 2006).

7.3. Effect on water use

Synonymous with the majority of abiotic perturbations, waterlogging tendsto alter nutrient and plant water use. Lowered metabolism and subsequentlyless available energy to drive nutrient transport across plasma membranesmay reduce nutrient absorption by roots (Cannell and Jackson, 1981; Drewand Stolzy, 1991). Water uptake can also decrease (Kramer, 1983 andreferences therein). Reductions in leaf gas exchange indicate the adverseeffect of waterlogging on plant water use (Davies et al., 2000b; Huang et al.,1994). Despite closure of the stomata, wilting and a decrease in leafC implythat the water transport from the root is insufficient to maintain the waterstatus of the shoot (Davies et al., 2000b).

Waterlogging and/or O2 deficiency tends to reduce Lpr. Kramer (1949,1983) reviewed the earlier literature and in the more recent literature theextent of the inhibition on Lpr appears to be species and treatment


dependent (Table 4). Apart from root death, Lpr may be influenced bywaterlogging in the longer term (days to weeks) through changes in rootstructure and anatomy (Section 3.2). For example, Erdmann et al. (1986)found that low O2 increased cell wall lignification and Huang et al. (1994)found that the xylem vessels of nodal roots decreased in diameter, althoughno measurements of root water transport were undertaken. Aerenchymaand barriers to ROL could also influence radial water flow, but this woulddepend on the length of the absorbing region. The formation of aeren-chyma and barriers to ROL do not appear to affect Lpr of rice andH. marinum roots (Garthwaite et al., 2006; Miyamoto et al., 2001;Ranathunge et al., 2003), but these measurements were undertaken inO2-sufficient conditions and water uptake in cereals occurs preferentiallyin the apical region (Section 3) where these anatomical features do notoccur.

A metabolically controlled mechanism could also influence Lpr underO2 deficiency, providing that a significant proportion of water flow acrossroots occurs via the cell-to-cell pathway, since respiration rates declinewhen the O2 supply is low. In further support of this, respiratory inhibitorshave been shown to reduce water flow through roots (Kamaluddin andZwiazek, 2001; Tyerman et al., 1992).

7.4. Anoxia and AQP activity

To date, no studies have directly investigated AQP transcript or proteinabundance (altered production or breakdown of in situ proteins) in plantroots in response to anoxia or hypoxia. However, several microarray ana-lyses monitored the expression of hundreds of A. thaliana genes, when theO2 concentration surrounding the roots or seedlings was reduced (Kloket al., 2002; Loreti et al., 2005; Liu et al., 2005). Cross-referencing thoselists of genes with GenBank (http://www.ncbi.nlm.nih.gov/Genbank/index.html; Benson et al., 2000) reveals that O2 deficiency modulates theexpression of some AQP genes (Table 5).

Klok et al. (2002) exposed cultured A. thaliana roots to hypoxia (5% O2

and 95% N2), which highly upregulated TIP1;2 in the first 0.5 h of treat-ment and TIP4;1 during 2–20 h of hypoxia. Submergence of the rootswill dilute the apoplast leading to uptake of water by cells and so TIPsmay be induced as a defence against adverse cytoplasmic volume changes(Section 5.3.1).

Several studies identified NIP genes that were rapidly and stronglyupregulated by anoxia or hypoxia (Table 5). Since these NIPs are aqua-glyceroporins, they may be induced to transport glycerol that accumulatesduring hypoxia (Weig et al., 2004). Additionally, NIPs could also beupregulated to facilitate the transport of gases.


Downregulation of AQP genes occurs later than upregulation, butoverall, hypoxia appears to repress water channel activity, especially after12 to 24 h of treatment (Liu et al., 2005; Table 5). Several of the PIPsdownregulated are predominantly expressed in roots (Table 2). Therefore,

Table 4 Summary of the effects of waterlogging or O2 deficiency on water transportin roots

Plant SpeciesWaterloggingtreatment

Reduction inLpr (or Lpc)(%) References

Agave deserti Anoxia (3 h) 0 Nobel et al., 1990

Arabidopsis thaliana Anoxia

(30 min)

42 Tournaire-Roux

et al., 2003

Helianthus annuus Anoxia

(16 h – 6 d)

46 (transient) Everard and Drew,

1989

Lupinus species Hypoxia

(30 min)

10 Bramley, 2006

Lupinus species

(cortical cells in

expanded region)

Hypoxia

(30 min)

70 Bramley, 2006

Lycopersicon

esculentum

Flooding

(8 h)

50 (transient) Bradford and Hsiao,

1982

Lycopersicon

esculentum

Flooding

(24 h)

0 Jackson et al., 1996

Populus tremuloides Hypoxia

(1–10 d)

26–75 Kamaluddin and

Zwiazek, 2002a

Ricinus communis Flooding

(2–24 h)

>50 Else et al., 2001

Triticum aestivum

(root cells near tip)

Hypoxia

(30 min)

85 Zhang and

Tyerman, 1991

Triticum aestivum

(cortical cells in

expanded region)

Hypoxia

(30 min)

46 Bramley, 2006

Triticum aestivum Hypoxia

(30 min)

20 Bramley, 2006

Zea mays Anoxia

(12 h)

28 Birner and Steudle,

1993

Zea mays Anoxia

(>20 h)

<70 Everard and Drew,

1987

Zea mays Hypoxia

(2 h)

27 (transient) Gibbs et al., 1998b

Zea mays Hypoxia

(30 min)

18 Tyerman et al., 1992

Zea mays (root cells) Hypoxia

(30 min)

60 Tyerman et al., 1992


Table

5AQPgeneexpressionin

roots

andwhole

seedlingsmodulatedbylowO2concentrations

Exp

ression

Loc

usa

Transcript

Details

Referen

ces

Upregulated

At3g26520

TIP1;2

Highlyupregulatedwithin

0.5

hofhypoxiatreatm

ent

Kloketal.,2002

At2g25810

TIP4;1

Veryhighlyupregulated2–20hofhypoxiatreatm

ent

Kloketal.,2002

At2g34390

NIP2;1

Rapidly(w

ithin

1h)andhighlyupregulatedin

response

to

anoxiaorhypoxia

Liu

etal.,2005;

Loretietal.,

2005

At2g29870

Pseudo-generapidly(w

ithin

1h)andstronglyupregulated

byhypoxia

Liu

etal.,2005

NIP

Highlyupregulatedin

thecentralcylinder

ofroots

Weigetal.,2004

AJ781013

NIP

Highlyupregulatedin

floodingtolerantrice

roots

Agarwaland

Grover,2005

At4g01470

TIP1;3

Transientlyupregulatedduring2–6hofhypoxia

Liu

etal.,2005

At1g31885

NIP3;1

Transientlyupregulatedduring6–12hhypoxia

Liu

etal.,2005

Downregulated

At3g61430

PIP1;1

Rapid

(2hhypoxia)andsustained

downregulation,but

expressionnotdownregulatedwhen

exogenoussucrose

applied

Liu

etal.,2005;

Loretietal.,

2005

At2g45960

PIP1;2

Sam

easPIP1;1

Liu

etal.,2005;

Loretietal.,

2005

At4g17340

TIP2;2

Sam

easPIP1;1

Liu

etal.,2005;

Loretietal.,

2005

(continued)

Table

5(continued)

Exp

ression

Locus

aTranscript

Details

References

At5g47450

TIP2;3

Sam

easPIP1;1,buteffectsofsucrose

notknown

Liu

etal.,2005

At2g39010

PIP2;6

Downregulatedwithadditionofexogenoussucrose

Loretietal.,

2005

At3g56950

SIP2;1

Sam

easPIP2;6

Loretietal.,

2005

At4g19030

NIP1;1

Sustained

downregulationafter12hhypoxia

Liu

etal.,2005

At2g45960

PIP1;2

Sam

easNIP1;1

Liu

etal.,2005

At1g01620

PIP1;3

Sam

easNIP1;1

Liu

etal.,2005

At3g53420

PIP2;1

Sam

easNIP1;1

Liu

etal.,2005

At2g37170

PIP2;2

Sam

easNIP1;1

Liu

etal.,2005

At5g60660

PIP2;4

Sam

easNIP1;1

Liu

etal.,2005

At4g35100

PIP2;7

Sam

easNIP1;1

Liu

etal.,2005

At3g26520

TIP1;2

Sam

easNIP1;1

Liu

etal.,2005

At3g16240

TIP2;1

Sam

easNIP1;1

Liu

etal.,2005

At4g23400

PIP1;5

Notdownregulateduntil24hofhypoxia

Liu

etal.,2005

At3g54820

PIP2;5

Sam

easPIP1;5

Liu

etal.,2005

At2g16850

PIP2;8

Sam

easPIP1;5

Liu

etal.,2005

At2g36830

TIP1;1

Sam

easPIP1;5

Liu

etal.,2005

aTheloci,identified

bymicroarrayanalysesofgenes

significantlyalteredbylow

O2,weregiven

inthesupplementaltablesattached

tothereferenceslisted

aboveand

wereusedto

search

thegeneticdatabaseGenBank(Bensonetal.2000)forthedescriptionoftherespectiveAQPtranscripts.

such a dramatic reduction in AQP gene transcripts could negatively impacton root water transport and potentially explain the increased root resistanceto water flow during waterlogging. Downregulation of TIPs and PIPs in theapical meristem could also be a component that reduces root growth duringwaterlogging.

Interestingly, Loreti et al. (2005) reported that the addition of exogenoussucrose altered which particular genes were downregulated by anoxia.Under anoxic conditions PIP1;1, PIP1;2 and TIP2;2 were downregulated,but not when sucrose was added to the bathing medium. conversely, PIP2;6and SIP2;1 were not downregulated by anoxia unless sucrose was present.Both of these latter genes are expressed in low amounts in roots and theproteins have unknown functions ( Jang et al., 2004; Quigley et al., 2001).The addition of sugars to the bathing medium often increases the toleranceof roots to anoxia, but by unknown mechanisms. Exogenous sucrose couldindirectly mitigate the reduction in Lpr induced by O2 deficiency.

Under field conditions, waterlogging is often transient, but no studieshave analyzed the expression of AQPs in roots after O2 deficiency followedby re-aeration. A problem with these studies could be separating the effectsof tissue death from adaptation under O2 deficiency. Few studies haveexamined AQP expression during a recovery period after other forms ofabiotic perturbations. However, Martre et al. (2002) found that PIPs areimportant in the recovery of A. thaliana after a water deficit. A key compo-nent in the recovery of a species after waterlogging could be the installationof new AQPs in root membranes, particularly if O2 deficiency leads to thebreakdown of AQPs or root death. Moreover, given that the turnover rateof some AQPs can occur within a few hours, the resumption of wateruptake and growth could also be rapid. After a short-term (0.5 h) mildhypoxia treatment followed by 1 h re-aeration, Lpr of wheat roots was�1.5times greater than initial values of Lpr, which was associated with an increasein AQP activity (Bramley, 2006). Moreover, in a long-term glasshouseexperiment waterlogged wheat roots rapidly resumed growth, when thepots were drained and the growth rate was greater than before the plantswere waterlogged (Bramley, 2006). An increase in AQP activity could notonly facilitate the rapid transport of water to the shoots, but may alsomediate changes in growth rate during the recovery period.

Rapid responses in Lpr to O2 deficiency may also be mediated by thegating of AQPs. As detailed in Section 5.3.3.1.2, protonation of PIPsduring cytosolic acidification closes the AQP pore and reduces the mem-brane permeability to water (Tournaire-Roux et al., 2003). Anoxia causesthe cytoplasm to become acidified (Menegus et al., 1991), which thereforemay close AQPs. The significance of this reaction on Lpr depends on thecontribution of AQPs to radial water flow, and also on the degree ofcytoplasmic acidification, which increases over time during anoxic condi-tions. However, some species such as rice are more tolerant of anoxia and


regulate their cytoplasmic pH to avoid acidosis (Menegus et al., 1991), but itis not known what effect this has on Lpc and hence on Lpr. Furthermore, innature plants experience a gradual decline in O2 rather than a suddenexposure to anaerobic conditions (i.e., hypoxia prior to anoxia), whichtriggers acclimatic responses such as avoidance of proton accumulation andexpression of proteins required for anaerobic respiration. A pretreatmentwith hypoxic conditions increases the tolerance of many species tosubsequent anoxia, and one of the processes related to this enhanced toler-ance is better regulation of cytoplasmic pH (Germain et al., 1997; Kato-Noguchi, 2000;Waters et al., 1991; Xia and Saglio, 1992).When exposed tohypoxic or anoxic conditions, the internal pH is generally not as low as thatused to demonstrate the inhibitory effects of pH on Lpc or Pf (Alleva et al.,2006; Menegus et al., 1991; Tournaire-Roux et al., 2003; Xia and Roberts,1994, 1996). However, the permeability of wheat and lupin rootcell membranes can decrease significantly, when exposed to only 0.5 h of3–4%O2 (Bramley, 2006; Zhang and Tyerman, 1991), which is considered arelatively mild hypoxic treatment. It is therefore, not clear whether a reduc-tion in internal pH is the initial factor that reduces Lpc through a closure ofAQPs. The effect may also be related to other mechanisms of AQP gatingsuch as changes in apoplastic water potential and osmoregulation that arisesfrom the effect of O2 deficiency on ion leakage from a depolarization of themembrane potential (Zhang and Tyerman, 1991, 1997). Calcium fluxes alsochange in response toO2 deficiency, which can affect AQP gating (Dat et al.,2004). Moreover, it has never been tested whether O2 itself affects AQPgating as could be demonstrated with stopflow spectrophotometry.

Overall, AQP activity does appear to decrease in response to O2 defi-ciency. This could have significant consequences on the rate of waterflow through roots, provided that a large proportion of water flow occursthrough the cell-to-cell pathway (Section 3.2.2.1). In addition, given thatsome AQPs such as PIP1s are permeable to other small neutral molecules, areduction in AQP activity may influence other cellular processes under O2

deficiency. For example, CO2 and ethanol build up under anaerobic con-ditions (Gibbs and Greenway, 2003), which may be because these mole-cules are permeable to some AQPs (Steudle and Henzler, 1995; Uehleinet al., 2003).

8. Conclusion

Water flow through roots can be controlled by a variety of mechan-isms. The structure of the root and the anatomy of the radial pathways forflow dominate the process because they determine the physical hydraulicconductance of the root system to water flow. Changes to this hydraulicconductance will be slow and growth dependent, as the root lays down new


tissue, extends and/or becomes suberized. AQPs can influence the rate ofwater flow through the radial pathway of roots, but only if the hydraulicconductance of the apoplast is less than that of the cell-to-cell pathway.AQPsmay increase the hydraulic conductivity of the cell-to-cell pathway byincreasing their abundance and/or opening their pores. The distinct advan-tage of a significant proportion of radial water flow through AQPs is that itprovides the ability for the plant to rapidly and reversibly regulate flow inresponse to a fluctuating environment. AQP transcripts and proteins havebeen found to vary significantly and rapidly in response to abiotic perturba-tions. Moreover, AQP gating provides an even finer control. Despite theabundance of AQP homologs in roots, indicating that they play a part inregulating root water flow, there are also many other important functionalroles that they may perform. The majority of research to date has focussedon the molecular aspects of AQP activity, but more detailed physiology isrequired to understand AQP functions.

Of those species where radial water flow occurs through the cell-to-cellpathway, AQPs may influence water transport through roots when plantsare waterlogged, since AQP genes tend to be downregulated underO2-deficient conditions. However, long-term exposure to O2 deficiencycauses root systems to die, particularly in less tolerant species, which hencewould reduce the absorbing surface area for water uptake. Root systems thatdo not die avoid anoxia by forming aerenchyma and barriers to ROL.These anatomical features may obstruct water flow through the radialpathway and measurements of their effect on Lpr have not been undertakenduring O2 deficiency. AQPs in the apical region are likely to be down-regulated and/or close as the cytoplasm becomes acidified, since this regionis the most sensitive to O2 deficiency. Conversely, AQP activity inO2-sufficient cells (e.g., adjacent arenchyma) could increase and hencecompensate for the reduced hydraulic conductance caused by anatomicalchanges. Although little has been investigated, AQPs are likely to bebeneficial during recovery when the root system is no longer submergedand the O2 levels in the rhizosphere return to normal. The speed in theresumption of growth and transport of water and nutrients are key featuresthat influence the tolerance of a species to waterlogging and all theseattributes can be influenced by AQP activity. For us to understand thephysiological mechanisms that enhance a species’ tolerance to waterlogging,it is important that more research focuses on AQP-mediated processes thatare affected by O2 deficiency.

ACKNOWLEDGMENTS

A scholarship for HB and funding for research were supported by the Grains ResearchDevelopment Corporation (GRDC) of Australia.


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