Role of aquaporins in leaf physiology

Journal of Experimental Botany, Vol. 60, No. 11, pp. 2971–2985, 2009 doi:10.1093/jxb/erp171 Advance Access publication 19 June, 2009

DARWIN REVIEW

Role of aquaporins in leaf physiology Robert B. Heinen*, Qing Ye* and Francxois Chaumont† Institut des Sciences de la Vie, Universite´ catholique de Louvain, Croix du Sud 5-15, B-1348 Louvain-la-Neuve, Belgium Received 2 February 2009; Revised 1 May 2009; Accepted 5 May 2009

Abstract

Key words: Aquaporin, carbon dioxide, growth, hydraulic conductance, leaf, water.

Introduction Plant growth and development are dependent on essential physiological processes taking place in the leaf, such as photosynthesis and transpiration. To function properly, the leaf needs to maintain a balanced water content. This is especially important during evapotranspiration, during which water is unavoidably lost at the leaf surface to allow CO2 uptake. Before evaporation through the stomata, the high amounts of water transported from the soil to the leaf (long-distance transport) must cross several cell layers of different tissues (radial water transport). As in the composite transport model in the root, water can take different paths on its way through the leaf, moving through the apoplast (cell walls) or from cell to cell (Steudle, 1994; Steudle and Peterson, 1998). The latter route is composed of the symplastic (through the plasmodesmata) and the transcellular (across cell membranes) paths. The contribution of each pathway to the overall leaf hydraulic conductance (Kleaf) is unclear. Due to its low resistance to water flux, the apoplastic path is believed to be the main route during

transpiration (Sack and Holbrook, 2006). However, several recent studies have attributed an important role to the cellto-cell path (Cochard et al., 2007; Kim and Steudle, 2007; Ye et al., 2008). The presence of lignified or suberized cell walls, which constitute apoplastic barriers, forces water to cross cell membranes, as do the Casparian strips in the root endodermis (Lersten, 1997; Fricke, 2002a; Hachez et al., 2008). In addition to radial water flux, which is controlled by evapotranspiration, water movement across the leaf cell membranes is also important for water homeostasis, in increasing cell volume, and in maintaining turgor during expansion, in regulating the opening and closure of stomata, and in controlling leaf movement. Water movement through cell membranes is facilitated by water channels, called aquaporins (AQPs). These proteins belong to the major intrinsic protein (MIP) family, members of which are found in almost all living organisms (Agre et al., 1998). AQPs have six membrane-spanning alpha helices and cytoplasmic N- and C- termini. The cytosolic

* These two authors contributed equally to this work. y To whom correspondence should be addressed: E-mail: [email protected] Abbreviations: AQP, aquaporin; CA, cold-acclimated; ER, endoplasmic reticulum; gs, stomatal conductance; gm, mesophyll conductance; gi, internal conductance; Kleaf, leaf hydraulic conductance; Lp, cell hydraulic conductivity; NA, non-acclimated; NIP, Nodulin26-like intrinsic protein; MIP, major intrinsic protein; PCMBS, p-chloromercuribenzenesulphonic acid; PIP, plasma membrane intrinsic protein; ROS, reactive oxygen species; SIP, small basic intrinsic protein; TIP, tonoplast intrinsic protein; WT, wild type; XIP, X intrinsic protein. ª The Author [2009]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved. For Permissions, please e-mail: [email protected]

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Playing a key role in plant growth and development, leaves need to be continuously supplied with water and carbon dioxide to fulfil their photosynthetic function. On its way through the leaf from the xylem to the stomata, water can either move through cell walls or pass from cell to cell to cross the different tissues. Although both pathways are probably used to some degree, evidence is accumulating that living cells contribute substantially to the overall leaf hydraulic conductance (Kleaf). Transcellular water flow is facilitated and regulated by water channels in the membranes, named aquaporins (AQPs). This review addresses how AQP expression and activity effectively regulate the leaf water balance in normal conditions and modify the cell membrane water permeability in response to different environmental factors, such as irradiance, temperature, and water supply. The role of AQPs in leaf growth and movement, and in CO2 transport is also discussed.

2972 | Heinen et al.

Leaf AQP expression in normal conditions In addition to gating and trafficking, expression is one of the most important means of AQP regulation. As AQPs

constitute a large and highly divergent protein family in plants (Chaumont et al., 2001; Johanson et al., 2001; Sakurai et al., 2005; Danielson and Johanson, 2008), the study of their expression level and localization is highly relevant to a better understanding of their diverse physiological roles.

Organ-specific expression Many studies comparing AQP expression in different plant organs might suggest that, under normal conditions, AQPs play a more important role in roots than leaves. Indeed, many PIP and TIP homologues studied have been reported to be expressed at lower levels in leaves than in roots (Johansson et al., 1996; Biela et al., 1999; Chaumont et al., 2000; Baiges et al., 2001; Smart et al., 2001; Zhang et al., 2008). For instance, Zea mays ZmPIP1;1, ZmPIP1;2, and ZmPIP2;5 transcript levels are higher in the roots compared to the leaves (Chaumont et al., 2000). This was confirmed using antibodies raised against amino-terminal peptides of several ZmPIPs (Hachez et al., 2006a, 2008). The Western blots showed higher amounts of ZmPIPs in roots than in leaves of 11-d-old plantlets or the 6th leaf of adult plants (Fig. 1). However, other studies have shown that some AQPs are expressed at the same amount, more abundantly or even exclusively in leaves compared to roots. A recent characterization of tulip PIPs showed that TgPIP2;1 and TgPIP2;2 are highly expressed in leaves, but only TgPIP2;2 transcripts are detected, at a low level, in roots (Azad et al., 2008). In Arabidopsis thaliana, AtPIP2;6 transcript levels measured by real-time PCR are higher in the aerial parts than the roots, while the expression of the other PIPs is either lower (AtPIP1;1, AtPIP2;2, AtPIP2;3) or not significantly different (Jang et al., 2004). In rice (Oryza sativa), the expression of most AQP genes quantified by reverse transcription-PCR shows clear organ specificity. The levels of mRNA for 14 AQP homologues are higher in leaf blades than roots, while only six are expressed at higher levels in roots than leaves (Sakurai et al., 2005, 2008). All these data suggest that AQPs play important roles, not only in root water relations but also in the physiology and development of leaves. However, to understand their involvement in leaf water and solute flow better, detailed knowledge of their expression at the cellular, rather than the organ level is required.

Expression according to developmental stage Various studies have shown that AQP expression in leaves is developmentally regulated. While some are highly expressed in young, elongating leaf tissues, others are expressed preferentially in fully developed, matured leaves (Sarda et al., 1997; Chaumont et al., 1998; Maestrini et al., 2004; Wei et al., 2007; Hachez et al., 2008). For example, in Z. mays leaves, the expression of all 13 ZmPIP genes except ZmPIP2;7 can be detected by real-time RT-PCR, although to different extents, and the expression of most isoforms depends on the developmental stage, with expression being


loop (loop B) between the second and third transmembrane domains and the extracellular loop (loop E) between the fifth and sixth transmembrane domains form short helices that are relatively hydrophobic and are inserted into the membrane from opposite sides. These two loops contain highly conserved Asn-Pro-Ala (NPA) motifs located in the middle of the pore (Murata et al., 2000). While some AQPs behave as ‘strict’ water channels, others can conduct a wide range of non-polar solutes, such as urea or glycerol, and the non-polar gas carbon dioxide, the polar gas ammonia, the reactive oxygen species (ROS) hydrogen peroxide, and the metalloids antimonite, arsenite, and silicon (Bienert et al., 2008; Maurel et al., 2008). AQPs are particularly abundant in higher plants. Based on DNA sequence similarities, they are classified into five different subfamilies, namely PIPs (plasma membrane intrinsic proteins), TIPs (tonoplast intrinsic proteins), NIPs (nodulin26-like intrinsic proteins), SIPs (small basic intrinsic proteins), and XIPs (X intrinsic proteins) (Chaumont et al., 2001; Johanson et al., 2001; Sakurai et al., 2005; Danielson and Johanson, 2008). Although water transport across membrane pores had been proposed previously (Dainty and Ginzburg, 1963; House, 1974; Finkelstein, 1987), the discovery of the molecular structure of AQPs and detailed studies of their function revolutionized studies on plant water relations, at least those involving membranes (Steudle and Henzler, 1995; Maurel, 1997; Kjellbom et al., 1999; Tyerman et al., 1999; Steudle, 2001). Evidence is accumulating that AQPs play an important role in plant hydraulic relations at the cell, tissue, organ, and whole plant level. They facilitate the rapid, passive exchange of water across cell membranes and are responsible for up to 95% of the water permeability of plasma membranes (Henzler and Steudle, 2004). In general, there are three ways by which water exchange across cell membranes is regulated by AQPs: (i) their level of expression, i.e. AQP abundance; (ii) their trafficking, i.e. after synthesis in the ER, AQPs have to move to different cellular locations via the secretory pathway; and (iii) channel gating, i.e. the open/closed status of AQPs. Many recent reviews have summarized the impressive amount of data on the molecular structure, function, and regulation of plant AQPs (Chaumont et al., 2005; Hachez et al., 2006a; Kaldenhoff and Fischer, 2006; Maurel, 2007; Kaldenhoff et al., 2008; Maurel et al., 2008). In this work, the focus is on plant leaf AQPs, as a number of recent papers have highlighted the importance of AQPs in leaf growth and function. Firstly, the expression data for leaf AQPs from different plant species that provide clues about their roles are summarized; secondly, the responses of leaf AQPs to multiple environmental factors are described; and, finally, the data on the involvement of AQPs in leaf growth, leaf movement, and CO2 uptake are discussed.

Aquaporins in leaves | 2973 cell-to-cell pathway where regulation of AQPs could be of great importance (Hachez et al., 2008). In addition, the high expression of AQPs in the transition zone between elongating and matured maize leaf is thought to be due to the development of the metaxylem (built-up of apoplastic barriers by lignification/suberization) and increasing radial water flow due to higher photosynthetic activity and transpiration of the exposed leaf (Hachez et al., 2008). Finally, the isoforms highly expressed in fully developed, matured leaves probably play various roles in physiological processes, such as phloem loading, xylem water exit, stomatal aperture, gas transport (e.g. CO2), and leaf movement.

Tissue-specific expression

highest at the end of the elongation zone (Hachez et al., 2008). In barley (Hordeum vulgare), HvPIP1;3 is highly expressed in mature leaf tissue and HvPIP1;5 and HvPIP1;6 in the elongation zone (Wei et al., 2007). These developmental-specific expression patterns presumably reflect the physiological roles of the different isoforms. Growth-related expression of some AQPs has been described in other organs, such as the hypocotyl or root, in various species (Ludevid et al., 1992; Chaumont et al., 1998; Higuchi et al., 1998; Eisenbarth and Weig, 2005). In addition to cell wall yielding and solute supply, tissue hydraulic conductance and, thus, AQP activity are thought to be potential factors limiting cell expansion and growth (Fricke, 2002a) (see also the section ‘AQPs and leaf growth’). As the elongation zone of the barley and maize leaf is always covered by the sheath of older leaves, cells of this tissue are never exposed to a dry atmosphere. Thus water movement within this tissue, which is mainly driven by gradients of osmotic potential, could dominantly use the


Fig. 1. Maize plasma membrane AQPs are more abundant in roots than in leaves. ZmPIP protein amounts in 11-d-old maize root and shoot, and leaf 6 of adult plant. Microsomal membranes were extracted and analysed by Western blotting using antibodies against ZmPIP1 (general), ZmPIP1;2, ZmPIP2 (general), ZmPIP2;1/ 2;2, ZmPIP2;5 and ZmPIP2;6 as described in Hachez et al. (2006a and 2008). A colloidal blue stained SDS-PAGE gel was used as a loading control. The position of the molecular mass markers is indicated.

To understand the role of individual AQP in leaves, information on tissue-specific expression is essential. Translational fusion, in situ hybridization, and immunolocalization are the most common approaches presently used to study the tissue-specific expression of a single isoform. Table 1 presents an overview of the tissue localization of AQPs in leaves of various plant species. The large variety of sites at which they are found reflects the wide range of tasks they have to perform. Indeed, AQP expression has been described in all leaf tissues in which high transmembrane water movement is required. However, since AQPs can facilitate the movement of other solutes or gas, such as CO2, their localization in specific cells might be related to functions other than hydraulic control. Plants have to face the dilemma of taking up CO2 while unavoidably losing water vapour through their stomata. Adapting to local or global changes, stomata are able to open and close rapidly as a result of osmosis-driven water movement between the guard cells and surrounding tissue (Hetherington and Woodward, 2003; Roelfsema and Hedrich, 2005). Guard cell-localized AQP expression has been detected for A. thaliana PIP1;2, sunflower (Helianthus annuus) TIP7 and TIP20, Nicotiana tabacum AQP1, Norway spruce (Picea abies) TIP, spinach (Spinacia oleracea) PIP1;1, broad bean (Vicia faba) PIP1, and maize ZmPIP1;2 and ZmPIP2;1/2;2 (Kaldenhoff et al., 1995; Sarda et al., 1997; Oliviusson et al., 2001; Sun et al., 2001; Cui et al., 2005; Fraysse et al., 2005; Hachez et al., 2008; Uehlein et al., 2008). In maize, ZmPIP2;1/2;2 and ZmPIP1;2 expression is not only detected in guard cells, but also in the adjacent subsidiary cells, a second cell type contributing to the physiology of the stomatal aperture (Hachez et al., 2008) (Fig. 2A, B). In sunflower, the increase in SunTIP7 transcript levels during stomatal closure suggests its involvement in this mechanism by increasing guard cell water efflux (Sarda et al., 1997). Recently, immunological and translational fusion approaches have shown that the CO2 diffusion-facilitating NtAQP1 is not only found in guard and mesophyll cell plasma membranes, but also in the inner chloroplast membranes, indicating that it plays a role in the CO2 assimilation process (Uehlein et al., 2008) (see also the section ‘Leaf AQPs and CO2 transport’). Detection of

2974 | Heinen et al. Table 1. Aquaporin tissue localization in leaves of various plant species Isoform

Leaf localization

Techniquea

Reference

Arabidopsis thaliana

PIP1;2 PIP1 MOD (PIP1) c-TIP PIP1 PIP TIP7 TIP20 PIP1;6 MipA (PIP1)

Guard cells, in/around vascular bundles Plasmalemmasomes of mesophyll cells Blade, hydatodes Bundle sheath > mesophyll > vascular parenchyma Bundle sheath > mesophyll > vascular parenchyma Mesophyll Guard cells Guard cells Epidermis and adjacent mesophyll Vascular tissue > mesophyll Phloem associated cells, mesophyll, minor veins Companion cells, sieve elements, young xylem vessels (low) Seedlings: apical meristem (high), petiole and stem: vascular tissues, glandular trichomes, subepidermal cells Stem: cells of the central part of xylem vessels (high), layer surrounding the central vascular structure Young developping xylem vessels, cells between xylem and phloem, phloem (low), mesophyll (low) Developing xylem, xylem parenchyma, companion cells, spongy parenchyma (especially around stomatal cavities) Plasma and inner chloroplast membrane of guard and mesophyll cells Mesophyll (high), vascular tissues (very low) Mesophyll (high), bundle sheath and epidermis (low) Mesophyll Mesophyll, epidermis, parenchyma Differentiating vascular tissues, guard cells, bundle sheath Trichomes, marginal meristem Vascular bundles, periderm, subperiderm Vascular bundles Vascular bundles, periderm, subperiderm Palisade and spongy parenchyma Phloem sieve elements Guard cells Guard cells Xylem parenchyma, companion cells, around phloem strands Leaf primordia Companion cells (high), xylem parenchyma (high), bundle sheath, blade mesophyll (low in seedlings), midrib parenchyma, guard cells, subsidiary cells (only mature plants), epidermis (only mature plants), apical meristem (low), leaf primordia Companion cells (high), xylem parenchyma (high), bundle sheath, blade mesophyll, midrib parenchyma, subsidiary cells, guard cells, epidermis, apical meristem, leaf primordia Companion cells (high), xylem parenchyma (high), bundle sheath, mesophyll, midrib parenchyma (low)

GUS, ISH, ISI ISI GUS ISI ISI TPI ISH ISH ISP ISH GUS (tobacco) ISI

Kaldenhoff et al., 1995 Robinson et al., 1996 Dixit et al., 2001 Frangne et al., 2001 Frangne et al., 2001 Ohshima et al., 2001 Sarda et al., 1997 Sarda et al., 1997 Wei et al., 2007 Yamada et al., 1995 Yamada and Bohnert, 2000 Kirch et al., 2000

GUS (tobacco)

Yamada et al., 1997

ISI

Kirch et al., 2000

ISI

Kirch et al., 2000

ISI

Otto et al., 2000

ISI, GFP

Uehlein et al., 2008

ISI ISI ISH ISH ISH

Sakurai et al., 2008 Sakurai et al., 2008 Li et al., 2008 Montalvo-Hernandez et al., 2008 Oliviusson et al., 2001

GUS (tobacco) TPI, ISI TPI, ISI TPI, ISI ISI ISI ISI ISH ISH ISH ISI

Jones and Mullet, 1995 Suga et al., 2003 Suga et al., 2003 Suga et al., 2003 Karlsson et al., 2000 Fraysse et al., 2005 Fraysse et al., 2005 Sun et al., 2001, Cui et al., 2005 Barrieu et al., 1998 Chaumont et al., 1998 Hachez et al., 2008

ISI

Hachez et al., 2008

ISI

Hachez et al., 2008

Brassica campestris Brassica napus Graptopetalum paraguayense Helianthus annuus Hordeum vulgare Mesembryanthemum crystallinum

MipB

MipF (TIP) Nicotiana tabacum

Oryza sativa

Phaseolus vulgaris (Carioca) Picea abies Pisum sativum Raphanus sativus

Spinacia oleracea

Vicia faba Zea mays

AQP1

PIP1 PIP2;1 PIP2;7 AQP1 (TIP) TIP Trg31 (PIP1) PIP1 PIP2;1 TIP1 d-TIP PIP1;2 PIP1;1 PIP1 TIP1 PIP1;2

PIP2;1/2;2

PIP2;5

a Techniques: GFP, promoter::GFP fusion; GUS, promoter::GUS fusion; ISH, in situ hybridization; ISI, in situ immunolocalization; ISP, in situ PCR; TPI, tissue print immunoblotting.

several AQPs in guard cells indicate that they contribute to the regulation of their volume and turgor. However, the way AQP activity is regulated by the environmental and internal signals is still unknown.

Besides their important role in CO2 assimilation and photosynthesis, mesophyll cells constitute an important element in the pathway taken by water in the transpiration stream. Whether water on its way to the substomatal cavity


Species

Aquaporins in leaves | 2975

takes the apoplastic path or the cell-to-cell path is still a matter of debate and probably differs according to the species and conditions (Sack and Holbrook, 2006; Cochard et al., 2007; Ye et al., 2008). An essential contribution of AQPs to the latter pathway seems likely (Fig. 2C), given their high expression in mesophyll cells of various plant species (Yamada et al., 1995; Robinson et al., 1996; Karlsson et al., 2000; Otto and Kaldenhoff, 2000; Yamada and Bohnert, 2000; Frangne et al., 2001; Ohshima et al., 2001; Hachez et al., 2008; Li et al., 2008; MontalvoHernandez et al., 2008; Sakurai et al., 2008; Uehlein et al., 2008). It should be noted that, in many species, part of the mesophyll volume is airspace with limited cell-to-cell contact (Sack and Holbrook, 2006). Thus, it cannot be excluded that AQPs in these cells may rather play a role in the osmotic adjustment of the cell water relations in response to changes of apoplastic water potential. NtAQP1 expression is particularly high in the mesophyll cells around stomatal cavities, possibly facilitating gas exchange (CO2 and water) between the plant and the atmosphere (Otto and Kaldenhoff, 2000; Uehlein et al., 2008). However, whether water evaporates in the close vicinity of the substomatal

cavity or deeper in the mesophyll tissue is still not clear (Boyer, 1985; Sack and Holbrook, 2006). Before entering the mesophyll, water has to cross the bundle sheath when flowing out of the vascular bundles. Due to its highly compacted cells and their sometimes lignified or suberized cell walls, the bundle sheath prevents exposure of the vascular tissues to the air in the intercellular spaces. In the case of the presence of apoplastic barriers, all substances including water leaving or entering the vascular bundle have to enter the symplast as in the root endodermis (Canny, 1990; Hachez et al., 2008; Obrien and Kuo, 1975). Water movement through these cell layers could therefore be efficiently controlled and enhanced by AQPs (Fig. 3B). This hypothesis is supported by the expression of several AQPs in bundle sheath cells or cells surrounding the vascular bundles, which has been reported for AtPIP1;2, Brassica napus c-TIP and PIP1, common ice plant (Mesembryanthemum crystallinum) MipB, rice OsPIP1, PaTIP, ZmPIP1;2, ZmPIP2;1/2;2, and ZmPIP2;5 (Kaldenhoff et al., 1995; Kirch et al., 2000; Frangne et al., 2001; Oliviusson et al., 2001; Hachez et al., 2008; Sakurai et al., 2008). Once in these cells, the water stream can continue


Fig. 2. Models of aquaporin-facilitated transmembrane water flow in Z. mays stomatal apparatus and mesophyll cells. (A) Schematic view on a stoma during its opening process. Maize stomata are composed of two guard cells (GC, light green) surrounded by two subsidiary cells (SC, light blue). During stomatal opening, solute accumulation in the guard cells decreases the osmotic potential. To reestablish the osmotic equilibrium, water follows the solutes into the guard cells inducing an increase in their volume and turgor pressure which finally causes the pore aperture allowing gas/vapour exchanges. The water influx could be facilitated by a high expression and activity of aquaporins in the guard cell plasma membrane but also in the surrounding subsidiary and epidermal (EP) cells. The latter cell types could play an important role in supplying the water required by the guard cells. (B) Schematic view of a stoma during its closure. During the closure, the decrease in water potential in the guard cells induces a rapid water efflux, which could also be facilitated by AQPs in a similar way as during the opening process. (C) Schematic view of a cross-section through a maize leaf showing the radial water flow during transpiration out of the bundle sheath towards the stomatal cavity. Water of the transpiration stream can follow either the apoplastic or the cell-to-cell pathway. The transcellular component of the cell-to-cell path could be increased by the presence of AQPs. Note that (i) AQP expression in mesophyll cells could also facilitate the CO2 uptake, which is not represented in this figure and (ii) plasmodesmata, important for the symplastic pathway, have been omitted to avoid an overcharge of the figure. All these models are deduced from immunocytochemistry data demonstrating the presence of ZmPIP proteins in the plasma membrane of these cells (Hachez et al., 2008). BS, bundle sheath; EP, epidermis; GC, guard cell; MP, mesophyll; SC, subsidiary cell.


symplastically or transcellularly, but the contribution of each pathway is unknown. Interestingly, when AQP expression is detected in a species’ leaves, it is often present in tissues of the vascular bundles (Table 1). For example, ZmTIP1, ZmPIP1;2, ZmPIP2;1/2;2, ZmPIP2;5, and NtAQP1 are highly expressed in xylem parenchyma (Barrieu et al., 1998; Otto and Kaldenhoff, 2000; Hachez et al., 2008), suggesting their involvement in water exit from the xylem vessels. Studies also indicate a role of xylem parenchyma located AQPs in repairing embolism (Holbrook and Zwieniecki, 1999;

Meinzer et al., 2001). The process involved in the refilling of gas-filled vessels to maintain the transpiration stream after cavitation is still unclear, but has been shown to be faster than expected, suggesting the involvement of water channels (Fig. 3C) (Meinzer et al., 2001). ZmTIP1, ZmPIP1;2, ZmPIP2;1/2;2, ZmPIP2;5, and NtAQP1 have also been shown to be highly abundant in phloem companion cells and this might be related to phloem sap loading (Fig. 3D). Moreover, McMipA and SoPIP1;2 are expressed in phloem sieve elements (Kirch et al., 2000; Fraysse et al., 2005). In species with apoplastic sugar


Fig. 3. Models of aquaporin-facilitated transmembrane water flow in Z. mays bundle sheath, xylem parenchyma, and phloem companion cells. (A) Cross-section through a midrib vascular bundle giving an overview of the location of the different sections shown in (B), (C), and (D). (B) Schematic view of a cross-section showing the bundle sheath (BS, light yellow) and radial water efflux out of the metaxylem (MX, blue). Before entering the mesophyll (MP), water has to cross the bundle sheath. The presence of apoplastic barriers (lignified/suberized cell walls, dark yellow) similar to Casparian strips in the root endodermis forces the water to cross the plasma membrane. Transmembrane water movement through the bundle sheath could therefore be efficiently controlled and enhanced by AQPs. (C) Schematic view of a longitudinal section through the metaxylem (MX) showing a gas embolism (GE) in the xylem vessel called cavitation. AQPs expressed in the xylem parenchyma (XP) could play an important role in the refilling process which is still not fully understood. (D) Schematic view of a cross-section through a phloem sieve element and its associated companion cell showing the potential implication of AQPs during apoplastic sugar loading. The C4 pathway of sucrose biosynthesis and phloem loading is represented by purple arrows starting in mesophyll cells with the first CO2 fixation, followed by the Calvin cycle in the bundle sheath cells, and finally leading to the sucrose synthesis (for a review, see Lunn and Furbank, 1999). On their way into the vascular bundle, the carbon assimilates follow the symplastic path through the plasmodesmata. Since sieve elements have no symplastic contact with surrounding cells except their associated companion cell (the few existing plasmodesmata are thought to be especially narrow or even sealed), apoplastic phloem-loading species need sucrose transporters to charge the carbohydrates into the phloem against their concentration gradient (Slewinski et al., 2009; Turgeon, 2006). The resulting decrease in the osmotic potential drives an equilibrating water influx. High AQP expression in companion cells might increase the membrane water permeability needed during phloem loading. All these models are deduced from immunocytochemistry data demonstrating the presence of ZmPIP proteins in the plasma membrane of these cells (Hachez et al., 2008). BS, bundle sheath; CC, companion cell; GE, gas embolism; MP, mesophyll; MX, metaxylem; PP, phloem parenchyma; SE, sieve element; XP, xylem parenchyma.


Effect of deregulation of plant AQP expression on leaf hydraulic parameters Altering the level of expression of a single AQP gene by reverse genetics, such as over-expression, silencing, and/or knockout, is an interesting strategy for elucidating the physiological function of specific AQP genes. Alteration of AQP expression by these approaches has been shown to affect leaf cell water permeability and, in some cases, other physiological parameters, such as water potential, water loss rate, and stomatal conductance, indicating the importance of AQP for leaf function. However it is difficult to generalize phenotypes at tissue and plant levels as different mechanisms are probably used by plants to compensate lower water permeability (Hachez et al., 2006b). In Arabidopsis plants with reduced expression of PIP1s and/or PIP2s, the hydraulic conductivity (Lp) of isolated leaf protoplasts is markedly reduced (Kaldenhoff et al., 1998; Martre et al., 2002). To compensate the reduced cell Lp, plants increase their root mass by 2.5–5 times as compared with wild type (WT) plants to ensure sufficient water supply to the plant. Overexpression of lily AqpL1 (a PIP1 gene) in tobacco greatly increases the osmotic water permeability of leaf protoplasts and the water conductivity of leaf cells (Ding et al., 2004). The transgenic plants consume more water than WT plants, have a higher stomatal density in their young leaves, and exhibit a higher stomatal aperture

induced by light. These data indicate that overexpression of this AQP gene, by enhancing the cell-to-cell pathway of water movement within the leaf, might act as an internal factor to adapt the stomatal apparatus and increase the leaf transpiration (Ding et al., 2004). Overexpression of B. napus BnPIP1 in tobacco also results in a faster swelling of leaf protoplasts in hypotonic medium (Yu et al., 2005). In the case of AtPIP1;2 overexpression in tobacco plants, a significant increase in plant growth rate, leaf transpiration rate, stomatal density, and photosynthetic efficiency is observed under favourable growing conditions (Aharon et al., 2003). In a drought-sensitive cultivar of rice, overexpression of OsPIP1;3 under the control of a stress-inducible promoter results in a higher leaf water potential and transpiration rate, indicating that OsPIP1;3 may play a role in drought resistance (Lian et al., 2004). Overexpression of barley HvPIP2;1 in rice leads to faster water loss by the leaf which may be due to an increased transpiration rate. Overexpressing lines also show an increased CO2 assimilation rate and a faster growth rate, possibly resulting from an increased internal CO2 conductance (see also the section ‘Leaf AQPs and CO2 transport’). However, the plants are more sensitive to water stress, a fact that may lead to the observed anatomical adaptations of the leaves such as smaller mesophyll cells and thicker leaf cell walls (Hanba et al., 2004). To be able to conclude about the function of a specific AQP by modifying its expression in plant, it is important to (i) characterize the transport specificity of the channel and (ii) carefully analyse possible compensation mechanisms by close homologues. For instance, expression of AQPs isolated from cucumber and figleaf gourd in Arabidopsis perturbs the expressions of endogenous AQPs, especially when plants are subjected to water stresses (Jang et al., 2007). It is therefore impossible to attribute the detected phenotypes to the transgene or to the altered expression of endogenous AQPs.

Responses of leaf AQPs to environmental conditions While many AQPs coexist in plants, most isoforms seem to have their own specific characteristics which allow the plant to develop and grow adequately in ever-changing external conditions.

Water stress Water deficit, initially sensed by the roots, can affect the regulation of water movement in the shoots. A number of studies have shown that water stresses, including drought and salinity, can alter leaf AQP expression and activity. In the aerial part of Arabidopsis plants, drought stress induced with 250 mM mannitol significantly alters the expression of most PIP genes (Jang et al., 2004). For example, drought treatment rapidly decreases levels of PIP1;5, PIP2;2, PIP2;3, and PIP2;6 transcripts to one-tenth of the levels in


loading, phloem sieve elements with their associated companion cells are symplastically isolated from the neighbouring tissues. Photoassimilates need to be taken from the apoplast by a companion cell-localized sucrose-H+ symporter (Giaquinta, 1983; Rae et al., 2005; Slewinski et al., 2009). The resulting decrease in the osmotic potential drives an equilibrating water influx. High AQP expression in companion cells might increase the membrane water permeability needed during phloem loading, an idea supported by inhibition of the process by exogenous application of the sulphhydryl-modifying compound pchloromercuribenzenesulphonic acid (PCMBS) (Bel et al., 1993). However, PCMBS is a non-specific inhibitor, affecting sucrose transporters, AQPs, and a wide range of other proteins. Specific AQP inhibitors are highly needed to further verify the role of AQPs in the above mentioned processes. AQP expression has also been detected in the epidermis/ periderm and trichomes (Jones and Mullet, 1995; Yamada et al., 1997; Suga et al., 2003; Wei et al., 2007; Hachez et al., 2008; Montalvo-Hernandez et al., 2008; Sakurai et al., 2008). Finally, several AQPs are expressed in leaf primordial cells and fast dividing apical or marginal meristem cells (Jones and Mullet, 1995; Yamada et al., 1997; Chaumont et al., 1998; Hachez et al., 2008). From all of these studies, it can be concluded that AQPs are present in all kind of tissues. However, while some isoforms are ubiquitously expressed under normal conditions, others are highly tissue-specific.


Freezing and cold stress Perennial plants are able to survive tough winters through a phenomenon called cold acclimation. Low non-freezing or subfreezing temperatures can increase the freezing tolerance of plants by 3–5 C. Northern blot analysis and comparison of expressed sequence tags from non-acclimated (NA) and

cold-acclimated (CA) Rhododendron catawbiense leaf tissues showed a 10-fold down-regulation of RcPIP2;1 in CA tissues (Wei et al., 2006). Similarly, overexpression of RcPIP2s and Panax ginseng PIP1 in transgenic Arabidopsis plants compromised their freezing tolerance and cold acclimation ability, which is presumably due to their decreased capacity to resist freeze desiccation (Peng et al., 2007, 2008). The authors further demonstrated that all 13 endogenous PIPs except PIP2;5 are down-regulated in wild-type Arabidopsis plants during cold acclimation, confirming previous data concerning the impact of different abiotic stresses on Arabidopsis PIP transcription in the aerial parts of 2-week-old seedlings (Jang et al., 2004). Similar down-regulation of most rice PIP genes was recorded when seedlings were chilled to 7 C, and their expression recovered on return to 28 C. During recovery, significantly higher expression of OsPIP1;1, OsPIP2;1, and OsPIP2;7 was seen in shoots of a chilling tolerant variety than in those from a chilling sensitive one (Yu et al., 2006). All these data suggest that down-regulation of PIP transcripts during cold acclimation helps to prevent freezeinduced cellular dehydration, leading to increased freezing tolerance. Moreover, rapid rehydration of the leaf after cold/freeze stress might be mediated by AQPs. However, these mechanisms might be species-dependent as, in wheat leaves, a large increase in PIP transcript levels was seen after cold acclimation (from 22 C to 4 C) (Herman et al., 2006).

Responses of leaf AQPs to irradiance In many plant species, the Kleaf can be increased several fold by high irradiance. The response of the Kleaf to light is rather fast, as shown in Quercus rubra, in which it increases after only 30 min of high irradiance (1000–1200 lmol m 2 s 1) (Tyree et al., 2005). The increase in the Kleaf probably involves AQPs, as it is inhibited by mercury compounds, which are well-known AQP inhibitors. The lightinduced increase in the Kleaf in the burr oak is reduced by HgCl2 treatment, and this effect can be reversed using 2mercaptoethanol (Voicu et al., 2008). Similar results were obtained in the sunflower, in which the increase in the Kleaf by light is sensitive to mercury inhibition (Nardini et al., 2005). Using a cell pressure probe, Kim and Steudle (2007) found, for the first time at the cellular level, that the membrane water permeability of individual parenchyma cells in the midrib of maize leaves increases by a factor of 3 after 30 min of illumination at intensities of 100— 200 lmol m 2 s 1. Interestingly, the same authors subsequently showed that higher light intensities (800 and 1800 lmol m 2 s 1) dramatically decrease the water permeability of these leaf cells, possibly due to oxidative gating of AQPs induced by high light (Kim and Steudle, 2009). All these studies suggest that the effect of light on leaf hydraulic properties is related to the regulation of AQPs. This idea is further supported by the results of an analysis of the expression of two PIP2 isoforms (JrPIP2;1 and JrPIP2;2) in walnut (Juglans regia) leaves which correlates with the leaf


normal condition. The expression of PIP1;1 initially increases, while that of PIP1;2, PIP2;7, and PIP2;8 remains constant during the first 12 h of mannitol treatment, then gradually decreases. Using a longer term (up to 12 d), but gradual, drought stress, Alexandersson et al. (2005) showed that PIP transcripts in Arabidopsis leaves are generally down-regulated, except those for AtPIP1;4 and AtPIP2;5, which are up-regulated. The different responses of AQP expression (up/down-regulation or no change) to water stresses suggest that AQP isoforms can be divided into different groups which contribute differently to water transport and regulation, with some being stress-responsive (Hachez et al., 2006b). Different responses of AQPs to water stress were found in upland (drought-resistant) and lowland (drought-sensitive) rice. PIP proteins increase markedly in the roots of both types, but only in upland rice leaves. OsPIP1;2, OsPIP1;3, OsPIP2;1, and OsPIP2;5 mRNA levels in roots and OsPIP1;2 and OsPIP1;3 mRNA levels in leaves are significantly up-regulated in upland rice, but their expression is unchanged or down-regulated in lowland rice (Lian et al., 2006), indicating that AQPs present in the same species, but in different cultivars, can respond differently to water stress depending on their tolerance to water deficits. Expression of the V. faba VfPIP1 gene in Arabidopsis improves drought resistance in the mutant plants, probably by promoting stomatal closure under drought conditions, thus highlighting an important role of AQPs when plants are subject to water stress (Cui et al., 2008). Using double antisense Arabidopsis plants with reduced amounts of both PIP1 and PIP2 proteins, Martre et al. (2002) found that the leaf hydraulic conductance was similar in the mutants and control plants. However, upon water stress application, the mutant plants recovered their hydraulic conductance and transpiration rates less rapidly than the control plants and had a significantly lower leaf water potential after rewatering. These data led the authors to conclude that PIPs play an important role in the recovery of Arabidopsis from water-deficient conditions. In the heavy-metal accumulator Brassica juncea, BjPIP1 expression is up-regulated in leaves subjected to drought, salt, or low temperature, or exposed to heavy metals (Zhang et al., 2008). When overexpressed in tobacco plants, BjPIP1 enhances drought and cadmium resistance by decreasing the transpiration rate and stomatal conductance, suggesting that it might increase abiotic stress resistance by maintaining a reasonable water status in tobacco leaves (Zhang et al., 2008). However, overexpression of AQPs is not always beneficial to the plants. For instance, tobacco plants overexpressing AtPIP1;2 wilt more rapidly than control plants under drought stress (Aharon et al., 2003).


Inhibition of AQPs by reactive oxygen species Among the many internal or external factors that can affect AQP activities, hydrogen peroxide (H2O2) has been shown to be an effective inhibitor of leaf AQPs. In Tradescantia fluminesis, H2O2 perfusion via the leaf petiole results in a drop in the water permeability of epidermal cells measured with a cell pressure probe (Ye et al., 2008). Similarly, H2O2 treatment following Fe2+ pretreatment decreases the water permeability of parenchyma cells in maize leaves by a factor of 30 (Kim and Steudle, 2008). The effect could be overcome using an antioxidant, suggesting that the inhibition of AQP activity is probably due to an oxidative gating by ROS, such as *OH radicals, which can be produced in the Fenton reaction (Ye and Steudle, 2006). However, on the basis of the lack of effect of H2O2 on the activity of individual AQPs expressed in Xenopus oocytes, Boursiac et al. (2008) proposed that ROS do not gate AQPs through a direct oxidative mechanism, but rather act through a cell signalling mechanism and showed that H2O2 induces internalization of plasma membrane AQPs in unidentified vesicles. This process could therefore decrease the cell membrane water permeability. In addition, recent studies showed that, instead of being inhibited by H2O2, several AQPs isoforms are able to facilitate H2O2 transport across the tonoplast and plasma membranes (Bienert et al., 2006, 2007; Dynowski et al., 2008). The exact means by which H2O2 regulates the cell membrane permeability is still under investigation, but could involve different mechanisms, including direct oxidative gating, signal transduction

leading to the internalization of AQP proteins, or modification of their phosphorylation status, a post-translational modification thought to regulate their gating (Johansson et al., 1998; Tornroth-Horsefield et al., 2006; Van Wilder et al., 2008). It has to be noted that ROS like H2O2 and *OH are very aggressive and reactive chemicals. It is possible that the inhibition of cell water permeability was also due to indirect effects on AQPs such as lipid peroxidation, which needs to be verified in future experiments. Nevertheless, responses of AQPs to ROS may represent an efficient way to adjust water relations and cope with different types of stresses from which plants may suffer.

AQPs and leaf growth Like other plant organs, leaves grow in size through the irreversible expansion of cells, a process that requires the continuous uptake of water. As mentioned above, AQPs are expressed in leaf elongation zones and it is therefore tempting to believe that they are involved in the growth process (Fricke and Chaumont, 2007). Evidence that AQPs are involved in leaf growth comes from cell water permeability measured from different zones of the leaf. Using a pressure probe, Volkov et al. (2007) found that the water permeability of barley epidermal cells in the leaf growing elongation zone is higher than that of cells in the emerged non-growing leaf zone. Similar results were obtained using an osmotic swelling assay with protoplasts of mesophyll cells sampled from the same zones (growing versus nongrowing) (Volkov et al., 2007). Barley HvPIP1;6, a functional water channel, was found to be specifically expressed in the leaf epidermis and its expression was the highest in the elongation zone, where it accounted for 85% of the expression of known barley PIP1s, suggesting that it could facilitate growth-associated water uptake into these cells (Wei et al., 2007). However, both salinity (100 mM NaCl) and nutrient source reduction (by removing the leaves supplying nutrients to the newly emerged leaf), which were shown previously to reduce the leaf elongation rate of the growing barley leaf (Fricke, 2002b; Fricke and Peters, 2002; Fricke et al., 2006), increase the expression of HvPIP1;6. These authors proposed that, if the increase in HvPIP1;6 water channel activity assisted leaf growth under stress conditions, it should occur primarily through posttranslational modifications, such as phosphorylation. Direct evidence for the important role of AQPs in leaf growth was provided by Uehlein et al. (2003). In tobacco, NtAQP1 overexpression increases leaf growth rate, an effect that can be attributed to NtAQP1 acting as both a water channel and a CO2 transporter (see also the section ‘Leaf AQPs and CO2 transport’). The process of leaf cell expansion is a complex process involving multiple cellular activities, amongst which modification of the membrane permeability through AQP regulation is probably one important factor. It would be interesting to measure growth rate of plants with altered expression of single AQP isoforms which were


hydraulic conductance, i.e. the increased Kleaf in response to light is associated with marked accumulation of PIP2 proteins, while both the Kleaf and levels of PIP2 decrease when the light is turned off (Cochard et al., 2007). This effect of light on leaf AQPs is reflected by a fluctuating expression pattern that follows the day/night cycle. In maize leaves, most of the ZmPIP genes are more highly expressed during the first hours of the light period than at the end of the day or at night (Hachez et al., 2008). This correlates with changes in the membrane water permeability measured using a cell pressure probe (C Hachez, W Fricke, F Chaumont, unpublished data). A diurnal oscillation in the expression of NtAQP1 was also found in tobacco leaf petioles, in which high expression is seen in the morning and low expression in the evening, and this is consistent with the diurnal changes in the osmotic water permeability of protoplasts isolated from the same part of the leaf (Siefritz et al., 2004). Similar results were obtained in motor cells of Samanean samam, in which expression of the AQP homologue gene SsAQP2 shows a diurnal rhythm concomitant with the diurnal regulation of water movement in these cells (Moshelion et al., 2002). The regulation of AQP expression and activity by irradiance seems important for leaf hydraulics. However, the mechanism(s) by which light modulates the Kleaf via AQP regulation is not yet fully understood, and more precise investigations on the relationships between light and leaf AQPs are needed.

2980 | Heinen et al. shown to be active in cell membrane water transport, and in parallel to determine hydraulic properties of these transgenic plants.

AQPs and leaf movement

Leaf AQPs and CO2 transport CO2 assimilation is one of the most important physiological functions of the leaf and is indispensable for the survival and development of the plant. In plants, gas exchange between the atmosphere and the site of carboxylation (CO2) or the site of evaporation (H2O) has often been described by Ohm’s law of serial resistances (Flexas et al., 2008; Sack and Holbrook, 2006). After crossing the boundary layer and the leaf surface via the stomata, atmospheric CO2 diffuses through the substomatal cavity and intercellular spaces. The cell wall, plasma membrane, and cytoplasm of the mesophyll cells constitute the next barriers on its way to the assimilation site in the chloroplasts. Diffusion through stomata is known as stomatal conductance (gs), while that between the internal cavities and the carboxylation site is called mesophyll conductance (gm) or internal conductance (gi) (for a review, see Flexas et al., 2008; Warren, 2008). Not only the gs, but also the gi can be sufficiently small to limit photosynthesis (Harley et al., 1992; Loreto et al., 1992; Bernacchi et al., 2002). Relative limitation by gi can account for up to 40% and be similar to that due to stomatal conductance (Warren, 2008). Recent research has indicated that AQPs could explain the rapid adaptation of the gi to changes in environmental conditions. Using the non-specific AQP blocker HgCl2, Terashima and Ono (2002) observed a significant decrease in the gi in V. faba and Phaseolus vulgaris leaves. Heterologous expression of human AQP1 in Xenopus oocytes results in a significant increase in the CO2 permeability of the oocyte membrane (Nakhoul et al., 1998). A few years later, tobacco NtAQP1 was shown to increase the CO2 permeability of oocyte membranes in a similar way (Uehlein et al., 2003). To monitor cellular CO2 transport, 14C incorporation in homologous plant systems was investigated using NtAQP1 silenced or overexpressing tobacco lines. While a lower NtAQP1 level resulted in reduced incorporation, increased NtAQP1 expression led to higher incorporation rates compared with controls. Moreover, net photosynthesis was reduced in the antisense lines and increased in the overexpressing lines (Uehlein et al., 2003). These results suggest that NtAQP1 also acts as a CO2 membrane-transport-facilitating protein


It is known that the diurnal and circadian folding and unfolding of tobacco leaves are determined by different growth rates of the upper and lower leaf surfaces. Siefritz et al. (2004) showed that NtAQP1 plays an essential role in leaf movements in tobacco: (i) immunodetection experiments demonstrated that the peaks of the oscillating expression pattern of NtAQP1 in leaf petioles coincide with leaf unfolding; (ii) using an osmotic swelling assay on protoplasts isolated from leaf petioles, it was shown that the cell membrane water permeability is high during the unfolding process (in the morning), and low in the evening; and (iii) mutated tobacco plants with down-regulated NtAQP1 expression show substantially lower cell water permeability than the control plants, which eventually leads to reduced leaf movement. In contrast to the irreversible process of leaf growth controlled by cell elongation, leaf movement can also be mediated by turgor-regulated reversible cell volume changes of motor organs called pulvini, which were described by Darwin in one of his publications more than a century ago (Darwin, 1880). This special organ can co-ordinate rapid water exchange across the cell membrane and induce swelling and shrinking of cells located on opposite sides of the tissue, resulting in the movement of leaves and leaflets. As mentioned previously, the water permeability of S. saman protoplasts isolated from pulvini motor cells was found to be regulated diurnally (Moshelion et al., 2002). Functional analyses in Xenopus oocytes showed that a PIP homologue gene (SsAQP2) isolated from these cells encodes an efficient water conducting channel (Moshelion et al., 2002). Interestingly, the SsAQP2 expression pattern shows diurnal and circadian oscillation, with the highest mRNA levels in the morning, probably to meet the requirement for high osmotic water fluxes across the cell membrane when the pulvini open. The contribution of the tonoplast AQP c-TIP to the fast movements of leaves and leaflets in Mimosa pudica was investigated (Fleurat-Lessard et al., 1997). Immunodetection experiments showed that this AQP is almost exclusively present in the large central vacuoles. It was proposed that c-TIP probably facilitates fast water efflux from motor cells, leading to simultaneous turgor loss and leaf movement. Another study revealed that the interaction of two plasma membrane AQPs (MpPIP1;1 and MpPIP2;1) from Mimosa pudica increases the oocyte membrane water permeability (Temmei et al., 2005). It is very likely that the interaction of these two proteins affects their trafficking, as demonstrated in maize (Zelazny et al., 2007). Furthermore, phosphorylation of a serine residue in MpPIP1;1 leads to an increase in the co-operative effect of the two isoforms on water channel activity (Temmei et al., 2005). The role of these mechanisms in controlling the cell

turgor pressure of motor cells remains to be demonstrated. Another example of leaf movement is the active trapping closure of the Venus flytrap (Dionaea muscipula) to capture insects. It is plausible that trap closure involves rapid water fluxes in the leaf cells which are driven by the cell osmotic pressure and are actively regulated by the plant (Brown, 1916; Forterre et al., 2005). Interestingly, AQP inhibitors decrease the speed of trap closure (Volkov et al., 2008). All these data indicate that AQPs represent important components involved in leaf movement in different species. The elucidation of the rapid signalling events and the exact molecular mechanisms leading to the regulation of water channel activity and cell turgor modification will be the next challenge.

Aquaporins in leaves | 2981 highly dynamic, regulated by environmental cues and affected by leaf developmental stages or anatomy (Sack and Holbrook, 2006). Methodology will need to be developed to quantify and compare the contribution of AQPs in the leaf water conductivity in various situations: for example, within a species exposed to different irradiance or drought conditions, in different plant species differing in anatomy (venation and mesophyll). Plant lines deregulated in AQP gene expression in specific leaf tissues or cells could be very useful to this purpose, but careful characterization of the compensation mechanisms including anatomical modifications or alteration of endogenous AQP gene expression will be required. Other physiological processes that have to be explored are leaf elongation, embolism recovery, and stomatal regulation. While the signal transduction pathway and mechanisms governing stomatal aperture and closure are well documented, the exact role and regulation of AQPs in these processes are still missing. Finally, the cellular mechanisms regulating AQP trafficking and gating and, therefore, their activity will need to be taken into account in these studies to get an overall picture of the role AQPs play in leaf physiology.

Acknowledgements This work was supported by grants from the Belgian National Fund for Scientific Research (FNRS), the Interuniversity Attraction Poles Programme—Belgian Science Policy, and the ‘Communaute´ francxaise de Belgique— Actions de Recherches Concerte´es’. QY was supported by an individual Marie Curie European fellowship. RH is a Research Fellow at the FNRS.

References Conclusions While the importance of AQPs in root water uptake has been long recognized, evidence for their physiological relevance at the leaf level and, more particularly, during evapotranspiration was only obtained recently. mRNA and protein expression in different cell types clearly indicate an important role of AQPs in different processes. Biophysical measurements enabled variations in membrane water permeability during growth and after environmental cues to be demonstrated. The first reverse genetics experiments provided evidence of the important roles of AQPs in water and CO2 movement at the cell and tissue levels. However, in-depth investigation of the function and regulation of specific aquaporin in leaf physiology is still required. It is striking to observe that, within a species, so many aquaporins are expressed in the leaf (Hachez et al., 2008). The transport specificity and especially CO2 conductance of each isoform have to be determined. The contribution of AQP-facilitated CO2 transport in leaf tissues of C3 and C4 plants needs be elucidated and integrated with the AQP water transport. The leaf hydraulic conductance is

Agre P, Bonhivers M, Borgnia MJ. 1998. The aquaporins, blueprints for cellular plumbing systems. Journal of Biological Chemistry 273, 14659–14662. Aharon R, Shahak Y, Wininger S, Bendov R, Kapulnik Y, Galili G. 2003. Overexpression of a plasma membrane aquaporin in transgenic tobacco improves plant vigor under favorable growth conditions but not under drought or salt stress. The Plant Cell 15, 439–447. Alexandersson E, Fraysse L, Sjovall-Larsen S, Gustavsson S, Fellert M, Karlsson M, Johanson U, Kjellbom P. 2005. Whole gene family expression and drought stress regulation of aquaporins. Plant Molecular Biology 59, 469–484. Azad AK, Katsuhara M, Sawa Y, Ishikawa T, Shibata H. 2008. Characterization of four plasma membrane aquaporins in tulip petals: a putative homolog is regulated by phosphorylation. Plant and Cell Physiology 49, 1196–1208. Baiges I, Schaffner AR, Mas A. 2001. Eight cDNA encoding putative aquaporins in Vitis hybrid Richter-110 and their differential expression. Journal of Experimental Botany 52, 1949–1951. Barrieu F, Chaumont F, Chrispeels MJ. 1998. High expression of the tonoplast aquaporin ZmTIP1 in epidermal and conducting tissues of maize. Plant Physiology 117, 1153–1163.


in planta. The first evidence for a direct link between AQPs and the gi was provided by Hanba et al. (2004). Overexpression of barley HvPIP2;1 in transgenic rice plants increases the gi in leaves by 40% compared to wild-type plants. However, the increase in internal CO2 conductance is accompanied by anatomical (stomatal density, mesophyll porosity, mesophyll cell wall thickness, etc) and physiological (Rubisco content) differences, which are known to have an impact on CO2 assimilation and internal conductance (Hanba et al., 2004). Such anatomical and physiological alterations were not induced in NtAQP1 antisense or overexpressing tobacco lines, in which differences in the gi were again linked to AQP expression levels (Flexas et al., 2007). In addition to being present in the plasma membrane, NtAQP1 was detected in the chloroplast inner membrane by immunological and green fluorescent proteinfusion approaches (Uehlein et al., 2008). Moreover, the CO2 permeability, analysed by stopped flow spectrophotometry, was significantly reduced in the chloroplast membrane vesicles of the NtAQP1 silenced plants compared to the wild type. All those studies suggest a direct involvement of AQPs in the internal CO2 conductance gi via modulation of the CO2 permeability of the cell membranes. However, while these data obtained in plants seem very convincing, the physiological relevance of AQP-facilitated transmembrane CO2 diffusion in mammalian cells remains a matter of debate and the assumption that the biomembrane is an effective barrier to CO2 diffusion has been called into question (Verkman, 2002). Recently, the influence of unstirred layers was investigated by microelectrode measurements on reconstituted planar lipid bilayers of different compositions (Missner et al., 2008). The results showed that transmembrane CO2 permeability is not limited by the membrane itself, but by unstirred layers.

2982 | Heinen et al. Bel AJE, Ammerlaan A, Dijk AA. 1993. A three-step screening procedure to identify the mode of phloem loading in intact leaves. Planta 192, 31–39.

Cui XH, Hao FS, Chen H, Chen J, Wang XC. 2008. Expression of the Vicia faba VfPIP1 gene in Arabidopsis thaliana plants improves their drought resistance. Journal of Plant Research 121, 207–214.

Bernacchi CJ, Portis AR, Nakano H, von Caemmerer S, Long SP. 2002. Temperature response of mesophyll conductance. implications for the determination of rubisco enzyme kinetics and for limitations to photosynthesis in vivo. Plant Physiology 130, 1992–1998.

Dainty J, Ginzburg BZ. 1963. Irreversible thermodynamics and frictional models of membrane processes, with particular reference to the cell membrane. Journal of Theoretical Biology 5, 256–265.

Biela A, Grote K, Otto B, Hoth S, Hedrich R, Kaldenhoff R. 1999. The Nicotiana tabacum plasma membrane aquaporin NtAQP1 is mercury-insensitive and permeable for glycerol. The Plant Journal 18, 565–570. Bienert GP, Moller ALB, Kristiansen KA, Schulz A, Moller IM, Schjoerring JK, Jahn TP. 2007. Specific aquaporins facilitate the diffusion of hydrogen peroxide across membranes. Journal of Biological Chemistry 282, 1183–1192.

Bienert GP, Schussler MD, Jahn TP. 2008. Metalloids: essential, beneficial or toxic? Major intrinsic proteins sort it out. Trends in Biochemical Sciences 33, 20–26. Boursiac Y, Boudet J, Postaire O, Luu DT, Tournaire-Roux C, Maurel C. 2008. Stimulus-induced down-regulation of root water transport involves reactive oxygen species-activated cell signalling and plasma membrane intrinsic protein internalization. The Plant Journal 56, 207–218. Boyer J. 1985. Water transport. Annual Review of Plant Physiology 36, 473–516. Brown WH. 1916. The mechanism of movement and the duration of the effect of stimulation in the leaves of Dionaea. American Journal of Botany 3, 68–90. Canny MJ. 1990. What becomes of the transpiration stream. New Phytologist 114, 341–368. Chaumont F, Barrieu F, Herman EM, Chrispeels MJ. 1998. Characterization of a maize tonoplast aquaporin expressed in zones of cell division and elongation. Plant Physiology 117, 1143–1152. Chaumont F, Barrieu F, Jung R, Chrispeels MJ. 2000. Plasma membrane intrinsic proteins from maize cluster in two sequence subgroups with differential aquaporin activity. Plant Physiology 122, 1025–1034. Chaumont F, Barrieu F, Wojcik E, Chrispeels MJ, Jung R. 2001. Aquaporins constitute a large and highly divergent protein family in maize. Plant Physiology 125, 1206–1215. Chaumont F, Moshelion M, Daniels MJ. 2005. Regulation of plant aquaporin activity. Biology of the Cell 97, 749–764.

Darwin C. 1880. The power of movement in plants. London: John Murray. Ding X, Iwasaki I, Kitagawa Y. 2004. Overexpression of a lily PIP1 gene in tobacco increased the osmotic water permeability of leaf cells. Plant, Cell and Environment 27, 177–186. Dixit R, Rizzo C, Nasrallah M, Nasrallah J. 2001. The brassica MIP-MOD gene encodes a functional water channel that is expressed in the stigma epidermis. Plant Molecular Biology 45, 51–62. Dynowski M, Schaaf G, Loque D, Moran O, Ludewig U. 2008. Plant plasma membrane water channels conduct the signalling molecule H2O2. Biochemical Journal 414, 53–61. Eisenbarth DA, Weig AR. 2005. Dynamics of aquaporins and water relations during hypocotyl elongation in Ricinus communis L. seedlings. Journal of Experimental Botany 56, 1831–1842. Finkelstein A. 1987. Water movement through lipid bilayers, pores and plasma membranes. New York: Wiley. Fleurat-Lessard P, Frangne N, Maeshima M, Ratajczak R, Bonnemain JL, Martinoia E. 1997. Increased expression of vacuolar aquaporin and H+-ATPase related to motor cell function in Mimosa pudica L. Plant Physiology 114, 827–834. Flexas J, Diaz-Espejo A, Galmes J, Kaldenhoff R, Medrano H, Ribas-Carbo M. 2007. Rapid variations of mesophyll conductance in response to changes in CO2 concentration around leaves. Plant, Cell and Environment 30, 1284–1298. Flexas J, Ribas-Carbo M, Diaz-Espej A, Galmes J, Medrano H. 2008. Mesophyll conductance to CO2: current knowledge and future prospects. Plant, Cell and Environment 31, 602–621. Forterre Y, Skotheim JM, Dumais J, Mahadevan L. 2005. How the Venus flytrap snaps. Nature 433, 421–425. Frangne N, Maeshima M, Schaffner AR, Mandel T, Martinoia E, Bonnemain JL. 2001. Expression and distribution of a vaculoar aquaporin in young and mature leaf tissues of Brassica napus in relation to water fluxes. Planta 212, 270–278. Fraysse LC, Wells B, McCann MC, Kjellbom P. 2005. Specific plasma membrane aquaporins of the PIP1 subfamily are expressed in sieve elements and guard cells. Biology of the Cell 97, 519–534.

Cochard H, Venisse J-S, Barigah TS, Brunel N, Herbette S, Guilliot A, Tyree MT, Sakr S. 2007. Putative role of aquaporins in variable hydraulic conductance of leaves in response to light. Plant Physiology 143, 122–133.

Fricke W. 2002a. Biophysical limitation of cell elongation in cereal leaves. Annals of Botany 90, 157–167.

Cui XH, Hao FS, Chen H, Cai JH, Chen J, Wang XC. 2005. Isolation and expression of an aquaporin-like gene VfPIP1 in Vicia faba. Progress in Natural Science 15, 496–501.

Fricke W, Akhiyarova G, Wei WX, et al. 2006. The short-term growth response to salt of the developing barley leaf. Journal of Experimental Botany 57, 1079–1095.

Fricke W. 2002b. Biophysical limitation of leaf cell elongation in source-reduced barley. Planta 215, 327–338.


Bienert GP, Schjoerring JK, Jahn TP. 2006. Membrane transport of hydrogen peroxide. Biochimica et Biophysica Acta, Biomembranes 1758, 994–1003.

Danielson JAH, Johanson U. 2008. Unexpected complexity of the aquaporin gene family in the moss Physcomitrella patens. BMC Plant Biology 8, 45.

Aquaporins in leaves | 2983 Fricke W, Chaumont F. 2007. Solute and water relations of growing plant cells. In: Verbelen JP, Vissenberg K, eds. The expanding cell, Vol. 5. Berlin, Heidelberg: Springer, 7–31.

expression patterns of endogenous aquaporin genes and alters plant responses to different stress conditions. Plant and Cell Physiology 48, 1331–1339.

Fricke W, Peters WS. 2002. The biophysics of leaf growth in saltstressed barley. A study at the cell level. Plant Physiology 129, 374–388. Giaquinta RT. 1983. Phloem loading of sucrose. Annual Review of Plant Physiology 34, 347–387.

Johanson U, Karlsson M, Johansson I, Gustavsson S, Sjovall S, Fraysse L, Weig AR, Kjellbom P. 2001. The complete set of genes encoding major intrinsic proteins in Arabidopsis provides a framework for a new nomenclature for major intrinsic proteins in plants. Plant Physiology 126, 1358–1369.

Hachez C, Heinen RB, Draye X, Chaumont F. 2008. The expression pattern of plasma membrane aquaporins in maize leaf highlights their role in hydraulic regulation. Plant Molecular Biology 68, 337–353.

Johansson I, Karlsson M, Shukla VK, Chrispeels MJ, Larsson C, Kjellbom P. 1998. Water transport activity of the plasma membrane aquaporin PM28A is regulated by phosphorylation. The Plant Cell 10, 451–459.

Hachez C, Moshelion M, Zelazny E, Cavez D, Chaumont F. 2006a. Localization and quantification of plasma membrane aquaporin expression in maize primary root: a clue to understanding their role as cellular plumbers. Plant Molecular Biology 62, 305–323.

Johansson I, Larsson C, Ek B, Kjellbom P. 1996. The major integral proteins of spinach leaf plasma membranes are putative aquaporins and are phosphorylated in response to Ca2+ and apoplastic water potential. The Plant Cell 8, 1181–1191.

Hanba YT, Shibasaka M, Hayashi Y, Hayakawa T, Kasamo K, Terashima I, Katsuhara M. 2004. Overexpression of the barley aquaporin HvPIP2;1 increases internal CO2 conductance and CO2 assimillation in the leaves of transgenic rice plants. Plant and Cell Physiology 45, 521–529. Harley PC, Loreto F, Dimarco G, Sharkey TD. 1992. Theoretical considerations when estimating the mesophyll conductance to CO2 flux by analysis of the response of photosynthesis to CO2. Plant Physiology 98, 1429–1436. Henzler T, Ye Q, Steudle E. 2004. Oxidative gating of water channels (aquaporins) in Chara by hydroxyl radicals. Plant, Cell and Environment 27, 1184–1195. Herman EM, Rotter K, Premakumar R, Elwinger G, Bae R, EhlerKing L, Chen SX, Livingston DP. 2006. Additional freeze hardiness in wheat acquired by exposure to –3 C is associated with extensive physiological, morphological, and molecular changes. Journal of Experimental Botany 57, 3601–3618. Hetherington AM, Woodward FI. 2003. The role of stomata in sensing and driving environmental change. Nature 424, 901–908. Higuchi T, Suga S, Tsuchiya T, Hisada H, Morishima S, Okada Y, Maeshima M. 1998. Molecular cloning, water channel activity and tissue specific expression of two isoforms of radish vacuolar aquaporin. Plant and Cell Physiology 39, 905–913. Holbrook NM, Zwieniecki MA. 1999. Embolism repair and xylem tension: do we need a miracle? Plant Physiology 120, 7–10. House CR. 1974. Water transport in cells and tissues. London: Edward Arnold. Jang JY, Kim DG, Kim YO, Kim JS, Kang H. 2004. An expression analysis of a gene family encoding plasma membrane aquaporins in response to abiotic stresses in Arabidopsis thaliana. Plant Molecular Biology 54, 713–725. Jang JY, Rhee JY, Kim DG, Chung GC, Lee JH, Kang H. 2007. Ectopic expression of a foreign aquaporin disrupts the natural

Kaldenhoff R, Fischer M. 2006. Aquaporins in plants. Acta Physiologica 187, 169–176. Kaldenhoff R, Grote K, Zhu JJ, Zimmermann U. 1998. Significance of plasmalemma aquaporins for water-transport in Arabidopsis thaliana. The Plant Journal 14, 121–128. Kaldenhoff R, Kolling A, Meyers J, Karmann U, Ruppel G, Richter G. 1995. The blue light-responsive AthH2 gene of Arabidopsis thaliana is primarily expressed in expanding as well as in differentiating cells and encodes a putative channel protein of the plasmalemma. The Plant Journal 7, 87–95. Kaldenhoff R, Ribas-Carbo M, Sans JF, Lovisolo C, Heckwolf M, Uehlein N. 2008. Aquaporins and plant water balance. Plant, Cell and Environment 31, 658–666. Karlsson M, Johansson I, Bush M, McCann MC, Maurel C, Larsson C, Kjellbom P. 2000. An abundant TIP expressed in mature highly vacuolated cells. The Plant Journal 21, 83–90. Kim YX, Steudle E. 2007. Light and turgor affect the water permeability (aquaporins) of parenchyma cells in the midrib of leaves of Zea mays. Journal of Experimental Botany 58, 4119–4129. Kim YX, Steudle E. 2009. Gating of aquaporins by light and reactive oxygen species in leaf parenchyma cells of the midrib of Zea mays. Journal of Experimental Botany 60, 547–556. Kirch HH, Vera-Estrella R, Golldack D, Quigley F, Michalowski CB, Barkla BJ, Bohnert HJ. 2000. Expression of water channel proteins in Mesembryanthemum crystallinum. Plant Physiology 123, 111–124. Kjellbom P, Larsson C, Johansson II , Karlsson M, Johanson U. 1999. Aquaporins and water homeostasis in plants. Trends in Plant Science 4, 308–314. Lersten NR. 1997. Occurrence of endodermis with a casparian strip in stem and leaf. Botanical Reviews 63, 265–272. Li GW, Zhang MH, Cai WM, Sun WN, Su WA. 2008. Characterization of OsPIP2;7, a water channel protein in rice. Plant and Cell Physiology 49, 1851–1858.


Hachez C, Zelazny E, Chaumont F. 2006b. Modulating the expression of aquaporin genes in planta: a key to understand their physiological functions? Biochimica et Biophysica Acta, Biomembranes 1758, 1142–1156.

Jones JT, Mullet JE. 1995. Developmental expression of a turgorresponsive gene that encodes an intrinsic membrane-protein. Plant Molecular Biology 28, 983–996.

2984 | Heinen et al. Lian HL, Yu X, Lane D, Sun WN, Tang ZC, Su WA. 2006. Upland rice and lowland rice exhibited different PIP expression under water deficit and ABA treatment. Cell Research 16, 651–660. Lian HL, Yu X, Ye Q, Ding X, Kitagawa Y, Kwak SS, Su WA, Tang ZC. 2004. The role of aquaporin RWC3 in drought avoidance in rice. Plant and Cell Physiology 45, 481–489. Loreto F, Harley PC, Dimarco G, Sharkey TD. 1992. Estimation of mesophyll conductance to CO2 flux by three different methods. Plant Physiology 98, 1437–1443. Ludevid D, Hofte H, Himelblau E, Chrispeels MJ. 1992. The expression pattern of the tonoplast intrinsic protein gamma-TIP in Arabidopsis thaliana is correlated with cell enlargement. Plant Physiology 100, 1633–1639. Lunn JE, Furbank RT. 1999. Sucrose biosynthesis in C-4 plants. New Phytologist 143, 221–237.

Martre P, Morillon R, Barrieu F, North GB, Nobel PS, Chrispeels MJ. 2002. Plasma membrane aquaporins play a significant role during recovery from water deficit. Plant Physiology 130, 2101–2110. Maurel C. 1997. Aquaporins and water permeability of plant membranes. Annual Review of Plant Physiology and Plant Molecular Biology 48, 399–429. Maurel C. 2007. Plant aquaporins: novel functions and regulation properties. FEBS Letters 581, 2227–2236. Maurel C, Verdoucq L, Luu DT, Santoni V. 2008. Plant aquaporins: membrane channels with multiple integrated functions. Annual Review of Plant Biology 59, 595–624. Meinzer FC, Clearwater MJ, Goldstein G. 2001. Water transport in trees: current perspectives, new insights and some controversies. Environmental and Experimental Botany 45, 239–262. Missner A, Kugler P, Saparov SM, Sommer K, Mathai JC, Zeidel ML, Pohl P. 2008. Carbon dioxide transport through membranes. Journal of Biological Chemistry 283, 25340–25347. Montalvo-Hernandez L, Piedra-Ibarra E, Gomez-Silva L, Lira-Carmona R, Acosta-Gallegos JA, Vazquez-Medrano J, Xoconostle-Cazares B, Ruiz-Medrano R. 2008. Differential accumulation of mRNAs in drought-tolerant and susceptible common bean cultivars in response to water deficit. New Phytologist 177, 102–113.

Obrien TP, Kuo J. 1975. Development of suberized lamella in mestome sheath of wheat leaves. Australian Journal of Botany 23, 783–794. Ohshima Y, Iwasaki I, Suga S, Murakami M, Inoue K, Maeshima M. 2001. Low aquaporin content and low osmotic water permeability of the plasma and vacuolar membranes of a CAM plant Graptopetalum paraguayense: comparison with radish. Plant and Cell Physiology 42, 1119–1129. Oliviusson P, Salaj J, Hakman I. 2001. Expression pattern of transcripts encoding water channel-like proteins in Norway spruce (Picea abies). Plant Molecular Biology 46, 289–299. Otto B, Kaldenhoff R. 2000. Cell-specific expression of the mercuryinsensitive plasma-membrane aquaporin NtAQP1 from Nicotiana tabacum. Planta 211, 167–172. Peng YH, Arora R, Li GW, Wang X, Fessehaie A. 2008. Rhododendron catawbiense plasma membrane intrinsic proteins are aquaporins, and their over-expression compromises constitutive freezing tolerance and cold acclimation ability of transgenic Arabidopsis plants. Plant, Cell and Environment 31, 1275–1289. Peng YH, Lin WL, Cai WM, Arora R. 2007. Overexpression of a Panax ginseng tonoplast aquaporin alters salt tolerance, drought tolerance and cold acclimation ability in transgenic Arabidopsis plants. Planta 226, 729–740. Rae AL, Perroux JM, Grof CPL. 2005. Sucrose partitioning between vascular bundles and storage parenchyma in the sugarcane stem: a potential role for the ShSUT1 sucrose transporter. Planta 220, 817–825. Robinson DG, Sieber H, Kammerloher W, Schaffner AR. 1996. PIP1 aquaporins are concentrated in plasmalemmasomes of Arabidopsis thaliana mesophyll. Plant Physiology 111, 645–649. Roelfsema MRG, Hedrich R. 2005. In the light of stomatal opening: new insights into ‘the Watergate’. New Phytologist 167, 665–691. Sack L, Holbrook NM. 2006. Leaf hydraulics. Annual Review of Plant Biology 57, 361–381. Sakurai J, Ahamed A, Murai M, Maeshima M, Uemura M. 2008. Tissue and cell-specific localization of rice aquaporins and their water transport activities. Plant and Cell Physiology 49, 30–39. Sakurai J, Ishikawa F, Yamaguchi T, Uemura M, Maeshima M. 2005. Identification of 33 rice aquaporin genes and analysis of their expression and function. Plant and Cell Physiology 46, 1568–1577.

Moshelion M, Becker D, Biela A, Uehlein N, Hedrich R, Otto B, Levi H, Moran N, Kaldenhoff R. 2002. Plasma membrane aquaporins in the motor cells of Samanea saman: diurnal and circadian regulation. The Plant Cell 14, 727–739.

Sarda X, Tousch D, Ferrare K, Legrand E, Dupuis JM, CasseDelbart F, Lamaze T. 1997. Two TIP-like genes encoding aquaporins are expressed in sunflower guard cells. The Plant Journal 12, 1103–1111.

Murata K, Mitsuoka K, Hirai T, Walz T, Agre P, Heymann JB, Engel A, Fujiyoshi Y. 2000. Structural determinants of water permeation through aquaporin-1. Nature 407, 599–605.

Siefritz F, Otto B, Bienert GP, van der Krol A, Kaldenhoff R. 2004. The plasma membrane aquaporin NtAQP1 is a key component of the leaf unfolding mechanism in tobacco. The Plant Journal 37, 147–155.

Nakhoul NL, Davis BA, Romero MF, Boron WF. 1998. Effect of expressing the water channel aquaporin-1 on the CO2 permeability of Xenopus oocytes. American Journal of Physiology, Cell Physiology 43, C543–C548.

Slewinski TL, Meeley R, Braun DM. 2009. Sucrose transporter1 functions in phloem loading in maize leaves. Journal of Experimental Botany 60, 881–892.


Maestrini P, Giordani T, Lunardi A, Cavallini A, Natali L. 2004. Isolation and expression of two aquaporin-encoding genes from the marine phanerogam Posidonia oceanica. Plant and Cell Physiology 45, 1838–1847.

Nardini A, Salleo S, Andri S. 2005. Circadian regulation of leaf hydraulic conductance in sunflower (Helianthus annuus cv. Margot). Plant, Cell and Environment 28, 750–759.

Aquaporins in leaves | 2985 Smart LB, Moskal WA, Cameron KD, Bennett AB. 2001. MIP genes are down-regulated under drought stress in Nicotiana glauca. Plant and Cell Physiology 42, 686–693.

Verkman AS. 2002. Does aquaporin-1 pass gas? An opposing view. Journal of Physiology 542, 31.

Steudle E. 1994. Water transport across roots. Plant and Soil 167, 79–90.

Voicu MC, Zwiazek JJ, Tyree MT. 2008. Light response of hydraulic conductance in bur oak (Quercus macrocarpa) leaves. Tree Physiology 28, 1007–1015.

Steudle E. 2001. The cohesion-tension mechanism and the acquisition of water by plant roots. Annual Review of Plant Physiology and Plant Molecular Biology 52, 847–875.

Volkov AG, Coopwood KJ, Markin VS. 2008. Inhibition of the Dionaea muscipula Ellis trap closure by ion and water channels blockers and uncouplers. Plant Science 175, 642–649.

Steudle E, Henzler T. 1995. Water channels in plants: do basic concepts of water transport change? Journal of Experimental Botany 46, 1067–1076.

Volkov V, Hachez C, Moshelion M, Draye X, Chaumont F, Fricke W. 2007. Water permeability differs between growing and nongrowing barley leaf tissues. Journal of Experimental Botany 58, 377–390.

Steudle E, Peterson CA. 1998. How does water get trough roots? Journal of Experimental Botany 49, 775–788.

Warren CR. 2008. Stand aside stomata, another actor deserves centre stage: the forgotten role of the internal conductance to CO2 transfer. Journal of Experimental Botany 59, 1475–1487.

Suga S, Murai M, Kuwagata T, Maeshima M. 2003. Differences in aquaporin levels among cell types of radish and measurement of osmotic water permeability of individual protoplasts. Plant and Cell Physiology 44, 277–286.

Temmei Y, Uchida S, Hoshino D, Kanzawa N, Kuwahara M, Sasaki S, Tsuchiya T. 2005. Water channel activities of Mimosa pudica plasma membrane intrinsic proteins are regulated by direct interaction and phosphorylation. FEBS Letters 579, 4417–4422. Terashima I, Ono K. 2002. Effects of HgCl2 on CO2 dependence of leaf photosynthesis: evidence indicating involvement of aquaporins in CO2 diffusion across the plasma membrane. Plant and Cell Physiology 43, 70–78. Tornroth-Horsefield S, Wang Y, Hedfalk K, Johanson U, Karlsson M, Tajkhorshid E, Neutze R, Kjellbom P. 2006. Structural mechanism of plant aquaporin gating. Nature 439, 688–694. Turgeon R. 2006. Phloem loading: how leaves gain their independence. Bioscience 56, 15–24. Tyerman SD, Bohnert HJ, Maurel C, Steudle E, Smith JAC. 1999. Plant aquaporins: their molecular biology, biophysics and significance for plant water relations. Journal of Experimental Botany 50, 1055–1071. Tyree MT, Nardini A, Salleo S, Sack L, El Omari B. 2005. The dependence of leaf hydraulic conductance on irradiance during HPFM measurements: any role for stomatal response? Journal of Experimental Botany 56, 737–744. Uehlein N, Lovisolo C, Siefritz F, Kaldenhoff R. 2003. The tobacco aquaporin NtAQP1 is a membrane CO2 pore with physiological functions. Nature 425, 734–737. Uehlein N, Otto B, Hanson DT, Fischer M, McDowell N, Kaldenhoff R. 2008. Function of Nicotiana tabacum aquaporins as chloroplast gas pores challenges the concept of membrane CO2 permeability. The Plant Cell 20, 648–657. Van Wilder V, Miecielica U, Degand H, Derua R, Waelkens E, Chaumont F. 2008. Maize plasma membrane aquaporins belonging to the PIP1 and PIP2 subgroups are in vivo phosphorylated. Plant and Cell Physiology 49, 1364–1377.

Wei W, Alexandersson E, Golldack D, Miller AJ, Kjellbom PO, Fricke W. 2007. HvPIP1;6, a barley (Hordeum vulgare L.) plasma membrane water channel particularly expressed in growing compared with non-growing leaf tissues. Plant and Cell Physiology 48, 1132–1147. Yamada S, Bohnert HJ. 2000. Expression of the PIP aquaporin promoter-MipA from the common ice plant in tobacco. Plant and Cell Physiology 41, 719–725. Yamada S, Katsuhara M, Kelly WB, Michalowski CB, Bohnert HJ. 1995. A family of transcripts encoding water channel proteins: tissue- specific expression in the common ice plant. The Plant Cell 7, 1129–1142. Yamada S, Nelson DE, Ley E, Marquez S, Bohnert HJ. 1997. The expression of an aquaporin promoter from Mesembryanthemum crystallinum in tobacco. Plant and Cell Physiology 38, 1326–1332. Ye Q, Holbrook NM, Zwieniecki MA. 2008. Cell-to-cell pathway dominates xylem-epidermis hydraulic connection in Tradescantia fluminensis (Vell. Conc.) leaves. Planta 227, 1311–1319. Ye Q, Steudle E. 2006. Oxidative gating of water channels (aquaporins) in corn roots. Plant, Cell and Environment 29, 459–470. Yu QJ, Hu YL, Li JF, Wu Q, Lin ZP. 2005. Sense and antisense expression of plasma membrane aquaporin BnPIP1 from Brassica napus in tobacco and its effects on plant drought resistance. Plant Science 169, 647–656. Yu X, Peng YH, Zhang MH, Shao YJ, Su WA, Tang ZC. 2006. Water relations and an expression analysis of plasma membrane intrinsic proteins in sensitive and tolerant rice during chilling and recovery. Cell Research 16, 599–608. Zelazny E, Borst JW, Muylaert M, Batoko H, Hemminga MA, Chaumont F. 2007. FRET imaging in living maize cells reveals that plasma membrane aquaporins interact to regulate their subcellular localization. Proceedings of the National Academy of Sciences, USA 104, 12359–12364. Zhang Y, Wang Z, Chai T, Wen Z, Zhang H. 2008. Indian mustard aquaporin improves drought and heavy-metal resistance in tobacco. Molecular Biotechnology 40, 280–292.


Sun MH, Xu W, Zhu YF, Su WA, Tang ZC. 2001. A simple method for in situ hybridization to RNA in guard cells of Vicia faba L.: the expression of aquaporins in guard cells. Plant Molecular Biology Reporter 19, 129–135.

Wei H, Dhanaraj AL, Arora R, Rowland LJ, Fu Y, Sun L. 2006. Identification of cold acclimation-responsive Rhododendron genes for lipid metabolism, membrane transport and lignin biosynthesis: importance of moderately abundant ESTs in genomic studies. Plant, Cell and Environment 29, 558–570.