Accepted Manuscript Title: CATION TRANSPORTERS/CHANNELS IN PLANTS: TOOLS FOR NUTRIENT BIOFORTIFICATION Author: Edgar Pinto Isabel Ferreira PII: DOI: Reference:
S0176-1617(15)00061-9 http://dx.doi.org/doi:10.1016/j.jplph.2015.02.010 JPLPH 52141
To appear in: Received date: Revised date: Accepted date:
15-12-2014 11-2-2015 11-2-2015
Please cite this article as: Pinto E, Ferreira I, CATION TRANSPORTERS/CHANNELS IN PLANTS: TOOLS FOR NUTRIENT BIOFORTIFICATION, Journal of Plant Physiology (2015), http://dx.doi.org/10.1016/j.jplph.2015.02.010 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
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CATION TRANSPORTERS/CHANNELS IN PLANTS:
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TOOLS FOR NUTRIENT BIOFORTIFICATION
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Edgar Pinto1, 2 *, Isabel Ferreira1
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REQUIMTE/ Department of Chemical Sciences, Laboratory of Bromatology and Hydrology, Faculty of Pharmacy - University of Porto, Portugal.
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CISA – Research Centre on Environment and Health, School of Allied Health
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Sciences, Polytechnic Institute of Porto, Portugal.
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Corresponding author address: Rua Jorge Viterbo Ferreira, 228, 4050-313 Porto,
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Portugal; email: [email protected]; Tel: +351220428642; Fax: +351226093390
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Abstract
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Cation transporters/channels are key players in a wide range of physiological functions
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in plants, including cell signaling, osmoregulation, plant nutrition and metal tolerance.
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The recent identification of genes encoding some of these transport systems has allowed
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new studies towards further understanding of their integrated roles in plant. This review
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summarizes recent discoveries regarding the function and regulation of the multiple
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systems involved in cation transport in plant cells. The role of membrane transport in
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the uptake, distribution and accumulation of cations in plant tissues, cell types and
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subcellular compartments is described. We also discuss how the knowledge of inter-
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and intra-species variation in cation uptake, transport and accumulation as well as the
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molecular mechanisms responsible for these processes can be used to increase nutrient
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phytoavailability and nutrients accumulation in the edible tissues of plants. The main
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trends for future research in the field of biofortification are proposed.
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Introduction
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Plant cells contain several cations. Potassium (K), calcium (Ca) and magnesium (Mg)
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are the major cationic species (Maathuis, 2009); iron (Fe), copper (Cu), manganese
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(Mn), nickel (Ni) and zinc (Zn) are widely recognized to be trace essential cations
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(Pilon et al., 2009) and cobalt (Co) and sodium (Na) are known as “beneficial” elements
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(Pilon-Smits et al., 2009).
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Each of the essential cations has a distinct specific role. Both Ca and Mg are
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constituents of organic structures, such as enzyme molecules, where they may be
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involved in the catalytic function of the enzymes (Maathuis, 2009). Potassium is one of
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the few cations that are not a component of organic structures. It is rather involved in
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the osmoregulation, the maintenance of electrochemical balance in cells and their
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compartments, and the regulation of the enzymatic activity (Wang and Wu, 2013). Due
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to their low concentration, trace essential cations do not play a direct role in
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osmoregulation or similar processes. These elements are redox-active and act as
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catalytically active cofactors in enzymes. Others have enzyme-activating functions, and
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others fulfil a structural role in stabilizing proteins (Hänsch and Mendel, 2009, Pilon-
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Smits et al., 2009).
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Cations must be acquired by plant roots from the soil solution and then delivered to
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tissues, organs and subcellular compartments, where they are required to support plants’
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metabolism (Maathuis, 2009). Plants face major variations in cation uptake, particularly
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in the case of trace essential (Fe, Zn, Cu, Ni and Mn) and beneficial (Co and Na)
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cations. On the one hand, the phytoavailability of these elements can be low, resulting
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in deficiency symptoms of the plants that grown in such environments. On the other
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hand, toxicity can occur in certain environments were the phytoavailability of
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problematic elements (e.g., Na) is enhanced by the conditions of that environment
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(Pinto et al., 2014b, Pinto et al., 2015). Plants have to cope with variable demands of
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different tissues during growth or in response to abiotic stresses. Thus, along the course
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of evolution, plants have developed highly specialized homeostatic mechanisms that
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deal with systemic spatiotemporal requirements in the acquisition, transport and supply
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of cations to specific tissues, organs or compartments (Pinto et al., 2014a).
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It is been widely recognized that the distinct patterns in the uptake, distribution and
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accumulation of cations in plants is the result of selective transport processes in which
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membrane transport proteins are involved (Haferkamp and Linka, 2012). The
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identification of several gene families encoding putative cation transporters/channels
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constituted a major breakthrough in the plant science field enabling the realization of
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studies at the gene and protein levels (e.g., gene expression at the tissue or plant levels,
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transport activity and subcellular localization) to achieve a further understanding of the
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integrated roles of these transport systems in plants.
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The list of essential minerals for plants is quite similar but not equal to the list for
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humans. For example, Se, I, Co and Na are essential for humans but not for most plants
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(White and Brown, 2010). Therefore, human deficiencies may arise for these elements
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in populations that depend on plants grown locally where the soil content of these
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elements is low. Actually, serious human deficiencies of Se and I exist worldwide
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(Hirschi, 2009). For nutrients required by both plants and humans, one can expect that
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human deficiencies will be less of a problem because the elements will necessarily be
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present in plant foods. However, the mineral content of plants is sometimes very low to
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meet human requirements, or the minerals may be present in forms that cannot be
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efficiently utilized by humans. Examples of both these situations are Ca and Fe,
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respectively (Jeong and Guerinot, 2008, Dayod et al., 2010). Thus, new strategies are
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needed to ensure a sustainable supply of healthy food to world’s population while
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meeting human nutritional requirements.
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The aim of this review is to join together information about the major cation
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transporters/channels involved in the uptake, distribution and accumulation of essential
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(K, Ca, Mg, Fe, Cu, Mn, Zn and Ni) and beneficial (Co and Na) elements and how this
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information can be used in biofortification programs. In the following sections, a
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detailed description of the transporters/channels function will be provided taking into
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consideration the four major processes that elements usually undergo – (1) uptake, (2)
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root compartmentation, (3) translocation and (4) leaf compartmentation. To simplify, in
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each of these four processes the transporters/channels involved in cation transport will
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be described according to the following division: (i) macroelements (K, Ca and Mg);
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(ii) trace essential elements (Fe, Cu, Mn, Ni and Zn); and (iii) beneficial elements (Co
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and Na). Then, a careful description of pathways to improve the biofortification of plant
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foods will be provided.
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Uptake
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Cation transport across the plant plasma membrane is mediated by an electrochemical
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gradient of protons generated by plasma membrane H+-ATPases. This “primary
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transport system” pumps protons out of the cell creating pH and electrical potential
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differences across the plasma membrane (Fox and Guerinot, 1998). At that point,
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“secondary transport systems” can utilize these gradients for many plant functions such
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as nutrients influx and efflux (Palmgren, 2001b). Cations enter the root either by
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crossing the plasma membrane of the root endodermal cells (symplastic transport) or by
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entering the root apoplast through the space between cells (apoplastic transport). They
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cannot pass through membranes without the aid of membrane transporter proteins.
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These transporter proteins occur naturally in several plant membranes (e.g., tonoplast,
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endoplasmic reticulum, mitochondria, chloroplasts) and, usually, multiple transporters
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exist for each element (Pinto et al., 2014a). For instance, rice (Oryza sativa) has at least
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seven Cu transporters (Yuan et al. , 2011). Each transporter has unique properties.
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When a low concentration of nutrients is present in the soil solution, their uptake
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usually requires the high-affinity transport system (HATS). By contrast, the low-affinity
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transport system (LATS) is used more when high concentrations of nutrients are
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present, such as in agricultural soils after fertilization. Furthermore, the abundance of
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each transporter varies with tissue type and environmental conditions, making the
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uptake of cations in plants a complex process (Karley and White, 2009, Puig and
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Penarrubia, 2009).
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Macroelements
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Potassium (K) is the most abundant cation in plant cells and plays a vital role in plant
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growth and development. Normally, K content in soil solution ranges between 1 and 10
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mM whereas intracellular K levels are maintained at 100 – 200 mM. Therefore, K
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uptake from the soil is obviously against a concentration gradient (Anschutz et al.,
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2014, Zörb et al., 2014). Similarly to most mineral nutrients, the main acquisition of K
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from the outside environment follows a biphasic form, defined as the sum of two
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distinguishable uptake mechanisms (HATS and LATS) at the root plasma membrane
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(Wang and Wu, 2013). Several genes appear to encode putative K transporters/channels
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in the plasma membrane of Arabidopsis root cells (Figure 1). They are grouped into
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four families, including HAK/KT/KUP (K+/H+ symporters), HKT/Trk (K+/Na+
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symporters), CHX (cation/H+ antiporters) and Shaker channels (Nieves-Cordones et al.,
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2014). The bulk of HATS that mediate the influx of K are catalyzed by members of the
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HAK/KT/KUP family (Very et al., 2014). In Arabidopsis, 10 of 13 AtKT/KUPs are
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expressed in root hairs. Five of them were expressed in root tip cells suggesting an
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essential role of these HATS in K uptake (Ahn et al., 2004). From these 5 AtKT/KUPs
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expressed in Arabidopsis roots, AtKUP1, AtKUP4 and AtHAK5 are transporters in the
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plasma membrane of root cells that seem to be engaged in the K influx (Ahn et al.,
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2004, Fulgenzi et al., 2008, Rubio et al., 2008). The physiological importance of
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HKT/Trk transporters for high-affinity K influx is not yet fully clear. Several plant
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studies have pointed out that most of HKT transporters function as Na transporters and
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only a limited number of these transporters are Na+/K+ symporters. Therefore, evidence
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supports a restricted role of the HKT family in K uptake (Very el al., 2014). Several
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members of the CHX family have been identified as K transporters. For example,
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AtCHX13 and AtCHX17 have been proposed as possible pathways for root K influx
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(Cellier et al., 2004, Zhao et al., 2008). Low-affinity K influx seems to be mediated by
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Shaker K-channels such as AtAKT1 and AtKC1, which are mainly expressed in root
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epidermal cells (Geiger et al., 2009, Ardie et al., 2010, Jeanguenin et al., 2011). Plant
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Shaker channels display extraordinary K selectivity. Consequently, the Shaker K-
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channels AtAKT1 and AtKC1 as well as the HAK/KT/KUP transporter AtHAK5 are
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responsible for almost all K influx (Nieves-Cordones et al., 2014).
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A peculiar electrochemical gradient exists for Ca in plant roots since cytosolic Ca levels
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are in the sub-µM range, while Ca concentrations in the soil solution are in the mM
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range (Karley and White, 2009). Calcium is taken up symplastically through
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nonselective cation channels (NSCCs) present in the root system. These can be divided
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according to their voltage dependence into depolarization-activated (DA-NSCCs),
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hyperpolarization-activated (HA-NSCCs) and voltage-insensitive (VI-NSCCs) types, as
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shown in Figure 1 (Demidchik and Maathuis, 2007). Regarding HA-NSCCs, members
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of the annexin gene family are likely to encode these Ca channels (Mortimer et al.,
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2008). For VI-NSCCs, two gene families, the plasma membrane-localized cyclic-
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nucleotide gated channels (CNGC) and the ionotropic glutamate receptor (GLR)
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homologues are proposed to mediate Ca uptake (Ma et al., 2009, Swarbreck et al., 2013,
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Forde, 2014). Despite the type of NSCC used to mediate Ca influx, the opening of these
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channels must be tightly regulated, because changes in cytosolic Ca levels trigger
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several specific responses to many environmental and developmental stimuli (McAinsh
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and Pittman, 2009).
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Magnesium concentration in soil solutions normally ranges from 8.5 to 125 mM.
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Cytosolic Mg levels are around 0.4 mM, thus Mg influx into root cells can be achieved
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by Mg-permeable cation channels (Karley and White, 2009). Despite that, members of
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the MRS2/MTG gene family expressed in Arabidopsis appear to dominate the Mg
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uptake across the root plasma membrane (Figure 1) (Gebert et al., 2009). A member of
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the A. thaliana Mg transport family, AtMGT1, was expressed in tobacco (Nicotiana
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benthamiana) and showed an increased Mg content (Deng et al., 2006). In addition to
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AtMGT1, other two MRS2/MTG members, AtMGT7 and AtMGT9, are expressed in
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Arabidopsis roots and have a key role in Mg uptake (Mao et al., 2008, Chen et al.,
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2009).
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Trace essential elements
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Several Fe species may be taken up by plant roots. The divalent form (Fe2+) is the main
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absorbed species but Fe3+ and Fe-chelates are also taken up. Plants have developed
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several mechanisms to enhance Fe uptake. Two main categories can be distinguished
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according to the plant species: Strategy I in non-graminaceous plants and Strategy II in
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graminaceous plants (Figure 2). The main processes of Strategy I (used by non-
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graminaceous plants such as A. thaliana) are the excretion of protons and phenolic
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compounds from the roots to the rhizosphere, reduction of ferric chelates at the root
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surface, and the absorption of the resulting ferrous ions across the root by plasma
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membrane transporters (Kobayashi and Nishizawa, 2012). The H+-ATPase (HA)
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transporters are responsible for the efflux of protons to promote the acidification of the
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rhizosphere (Santi and Schmidt, 2009) and the recently identified OsPEZ1 and OsPEZ2
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(phenolics efflux zero) seems to be involved in the secretion of phenolics (Bashir et al.,
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2011a, Ishimaru et al., 2011). Non-graminaceous plants depend on ferric reductase
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oxidase (e.g., AtFRO2) to reduce the ferric chelates at the root surface (Jeong and
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Connolly, 2009) and then on the iron-regulated transporter (AtIRT1, AtIRT2 and
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AtIRT3) to absorb the generated Fe2+ ions across the root plasma membrane (Lin et al.,
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2009, Vert et al., 2009, Barberon et al., 2011). The Strategy II, used by graminaceous
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plants (e.g., barley, maize and rice), depends on the biosynthesis and secretion of
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phytosiderophores (PS) such as mugineic acids (MAs) which mobilize Fe (Kobayashi
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and Nishizawa, 2012). Recently, the gene responsible for MA secretion was identified
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to be the transporter of the mugineic acid family phytosiderophores 1 (OsTOM1 and
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HvTOM1, for rice and barley, respectively) (Nozoye et al., 2011). The MAs secreted
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into the rhizosphere stimulate the solubilization of Fe3+ in the soil and the resulting Fe-
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PS complexes can be taken up through Yellow Sripe (YS) and Yellow Stripe-like (YSL)
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transporters such as HvYS1, ZmYS1 and OsYSL15 (Murata et al., 2008, Inoue et al.,
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2009, Ueno et al., 2009, Nozoye et al., 2013). Despite being Strategy II plants,
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graminaceous plants (e.g., rice), can also have ferrous transporters (e.g., OsIRT1,
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OsNRAMP1 and OsNRAMP5) allowing the absorption of Fe2+ in addition to its
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Strategy II–based Fe3+-MA uptake (Takahashi et al., 2011, Ishimaru et al., 2012).
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The most bioavailable form of Cu in soils is Cu2+. However, root uptake is often
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facilitated by reduction of Cu2+ to Cu+. The ferric reductase oxidases AtFRO2, AtFRO3,
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AtFRO4 and AtFRO5, expressed in A. thaliana, seem to be involved in this process
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(Mukherjee et al., 2006, Bernal et al., 2012). Copper is likely to enter the cytosol of root
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cells through a cell surface COPT/Ctr-family transporter. Six members of the Ctr family
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(COPT1-6) which mediate the influx of Cu have been identified in Arabidopsis.
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AtCOPT1, AtCOPT2 and AtCOPT3 are expressed in the plasma membrane of roots and
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have an important role in the acquisition of Cu monovalent ions (Figure 3) (Yamasaki et
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al., 2009, Andres-Colas et al., 2010). Furthermore, members of the Zinc-Regulated
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Transporter and Iron-Regulated Transporter (ZRT-IRT)-like proteins (ZIP) family can
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also mediate the uptake and transport of Cu in Arabidopsis (Yamasaki et al., 2009, del
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Pozo et al., 2010). Members of the YS and YSL family (e.g., ZmYS1 and HvYSL2) are
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also involved in the transport of Cu-PS complexes (Murata et al., 2008, Araki et al.,
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2011).
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The oxidized forms Mn3+ and Mn4+ are not bioavailable to plants. It is the reduced form
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of this element, Mn2+, that is absorbed by root cells (Pittman, 2005). The plasma
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membrane IRT1, expressed in both Arabidopsis and barley, can transport Mn (Figure 3)
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(Pedas et al., 2008, Barberon et al., 2011). The overexpression of AtIRT2 leads to
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overaccumulation of Mn in transgenic plants, but the role of this transporter in Mn
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uptake remains to be clarified (Vert et al., 2009). The plasma membrane-localized
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NRAMP1 and NRAMP5 were shown to be high-affinity Mn transporters in Arabidopsis
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and rice, respectively. Both AtNRAMP1 and OsNRAMP5 have broad selectivity, and
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their expression is restricted to the root (Cailliatte et al., 2010, Ishimaru et al., 2012). In
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maize and barley, Mn uptake can also be performed by the ZmYS1 and HvYSL2
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transporters, respectively, which were confirmed to be capable of acquiring Mn-PS
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complexes from the rhizosphere (Murata et al., 2008, Araki et al., 2011).
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Nickel is present in the environment usually in the form of Ni2+, which is the most
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available form to plants. Divalent cations such as Ni, Cu and Fe have very similar
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physicochemical properties thus they compete with each other in biochemical and
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physiological processes in plants (Ghasemi et al., 2009). Nickel can enter symplastically
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into the root cells through AtIRT1 (Nishida et al., 2012). Moreover, Ni can also be
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taken up in the form of Ni-PS complexes by the ZmYS1 due to its broad selectivity for
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several cations (Figure 3) (Murata et al., 2008).
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Zinc is taken up across the plasma membrane of root cells as a free ion and/or
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complexed with PS (Sinclair and Kramer, 2012). The ZIP transporter family includes
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the best candidates for facilitating Zn influx into the plant cytoplasm. In addition to its
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role in Fe acquisition, the broad selectivity of AtIRT1 allows this transporter to mediate
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also the uptake of several divalent metal cations, including Zn (Figure 3) (Barberon et
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al., 2011, Fukao et al., 2011, Shanmugam et al., 2011). This was also demonstrated in
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rice plants overexpressing OsIRT1, which accumulate elevated levels of Zn in the
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shoots, roots and mature seeds (Lee and An, 2009). Like their close homologue AtIRT1,
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AtIRT2 and AtIRT3 are also able to transport Zn into roots (Lin et al., 2009, Vert et al.,
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2009). Other ZIP transporters are involved in the uptake of Zn from soil. For example,
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OsZIP1 and OsZIP3 are likely to play a role in this process (Bashir et al., 2012). In
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Arabidopsis, AtZIP4 expression is induced upon Zn deficiency, showing its function as
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a Zn transporter (Assunção et al., 2010). The rice plasma membrane-localized OsZIP8
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also seems to be a Zn transporter that functions in Zn uptake (Lee et al., 2010). A
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member of the NRAMP family, AtNRAMP1, is also able to mediate Zn influx into the
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root cells of Arabidopsis (Cailliatte et al., 2010). Zinc is also taken up from the
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rhizosphere in the form of Zn–PS complexes by members of the YS and YSL family
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(Murata et al., 2008, Araki et al., 2011).
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Beneficial elements
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Since soil salinity is a global environmental challenge and NaCl is typically the major
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salt that contributes to that salinity, much research activity has been dedicated to the
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characterization of Na transport and distribution in plants, particularly to its uptake by
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plant roots (Zhang et al., 2010). Sodium can enter the root cells by NSCCs, more
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specifically by the VI-NSCCs. In fact, VI-NSCC permeability to Na has been
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demonstrated in several tissues and species, resulting in substantial evidence that root
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Na influx is mediated by these channels (Demidchik and Maathuis, 2007). Another
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candidate for mediating Na influx is the low-affinity cation transporter (LCT) 1.
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However, this function is still not clear and further investigation must be carried out
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(Amtmann et al., 2001). K and Na ions compete with each other to enter plant cells due
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to their physicochemical similarity. Therefore, it is plausible that K transporters may
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also be involved in Na acquisition and distribution. However, the involvement of K
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transporters from both the HAK/KT/KUP and AKT families in Na movement is
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somewhat contradictory. Some studies support the idea that some members of the
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HAK/KT/KUP and AKT families are involved in Na accumulation (Ardie et al., 2010,
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Horie et al., 2011, Zhang et al., 2013) but others fail to confer a specific role to these
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transporters in Na influx (Nieves-Cordones et al., 2010). Significant evidence exists for
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the involvement of K transporters from the HKT families in the Na influx (Benito et al.,
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2014). The HKT family can be divided in two different sub-families according to their
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properties regarding Na and K transport in heterologous expression systems. Sub-family
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1, which shows Na-specific transport activity and mediates Na uptake, includes, among
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others, AtHKT1;1 and its orthologues OsHKT1;1 and HvHKT1;1 expressed in rice and
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barley, respectively (Haro et al., 2005, Sunarpi et al., 2005, Davenport et al., 2007,
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Jabnoune et al., 2009). Sub-family 2 functions as a K-Na co-transporter and includes,
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among others, HvHKT2;1, OsHKT2;1, OsHKT2;2 and OsHKT2;4 (Yao et al., 2010,
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Mian et al., 2011, Sassi et al., 2012). It should be noted that the involvement of a certain
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transporter may be genotype-specific and not transversal to all plant species.
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The mechanisms of Co uptake remain poorly understood, although some transporters
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have been identified. Cobalt can be taken up in its ionic form Co2+ through the plasma
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membrane-localized AtIRT1 (Vert et al., 2002) and in the form of Co-PS complex by
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the ZmYS1 in maize (Murata et al., 2008). However, other membrane transporter,
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AtNRAMP1, shows broad selectivity for divalent cations and can also participate in the
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influx of Co into root cells (Cailliatte et al., 2010).
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Root compartmentation
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Once inside plant root cells, cations can be stored in vacuoles, transformed and/or
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translocated to shoot tissues. Evidence suggests that monovalent cations have a crucial
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role in the osmotic potential of cells. By contrast, divalent cations, like Ca 2+, are usually
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complexed with organic compounds within cells and thus, are responsible for other
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tasks inside plant tissues (Martinoia et al., 2012). Plants have developed effective
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mechanisms to perform the chelation and sequestration of cations, such as induction of
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higher levels of chelating species, compartmentation in vacuoles and adsorption to the
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cell wall (Figure 4). Nicotianamine (NA), glutathione (GSH), phytochelatins (PCs),
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metallothioneins (MTs), and organic acids have been increasingly shown in the
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literature to play a key role in the chelation and sequestration of several cations (Guo et
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al., 2008, Larbi et al., 2010, Pal and Rai, 2010, Deinlein et al., 2012, Haydon et al.,
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2012).
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With respect to root compartmentation, cations may be stored in the vacuole and/or cell
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wall. The vacuole is generally considered to be the main storage site for cations in plant
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cells and vacuolar compartmentation of elements is also a part of the tolerance
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mechanism (Martinoia et al., 2012). However, cell wall proteins and pectins are also
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involved in metal chelation and tolerance (Larras et al., 2013).
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Macroelements
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In addition to its role in the maintenance of cell osmoticum, K is also required in several
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biochemical reactions. Since K is less chaotropic than Na, K is preferentially
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accumulated in plant cells (Wang and Wu, 2013). Once in the cytosol, K can cross the
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tonoplast through transporters or channels. Despite the limited knowledge of the
312
mechanisms that regulate K movement across the tonoplast, evidence suggests that
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transporters are responsible for the influx of K into vacuoles while channels mainly
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mediate its efflux to the cytosol (Martinoia et al., 2012, Ahmad and Maathuis, 2014).
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The Na+/H+ antiporters (NHX) are a class of transporters that catalyzes the
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electroneutral exchange of Na and/or K for H+ by using the electrochemical H+ gradient
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produced by the H+–ATPases present in the plasma membrane of cell organelles
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(Ahmad and Maathuis, 2014, Pottosin and Dobrovinskaya, 2014). In Arabidopsis
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thaliana, six intracellular NHX members are identified and divided into two groups,
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vacuolar (NHX1 to NHX4) and endosomal (NHX5 and NHX6) (Bassil et al., 2011).
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From the four vacuolar NHX transporters, two (NHX1 and NHX2) have been fully
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characterized and have a crucial role in the regulation of intra-vacuolar K in root cells
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(Barragan et al., 2012). Regarding K efflux to the cytosol, two families of K-permeable
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channels, TPC1 (two-pore channel1) and the TPK/KCOs (two-pore K channels), have
325
been identified in the tonoplast of root cells (Martinoia et al., 2012). The first class of K
326
channels is the TPC1 which was the first vacuolar channel to be identified and mediates
327
the Ca-dependent K, Na and Mg efflux from the vacuole (Hedrich and Marten, 2011). It
328
has been suggested a possible role of TPC1 in Ca efflux from the vacuoles but some
329
contradictory data about this issue exist in the literature (Ranf et al., 2008). The
330
TPK/KCO family constitutes the second class of K channels located in the vacuolar
331
membrane. In Arabidopsis thaliana, the AtTPK/KCO family comprises five two-pore K
332
channels (TPKs) and one Kir-type K channels (KCO). From the five TPKs, four
333
(AtTPK1, -2, -3, and -5) are localized in the tonoplast as well as the AtKCO3 (Voelker
334
et al., 2010). Characterization of Arabidopsis tpk1 knockout mutants and TPK1-
335
overexpressing lines demonstrate that TPK1 has an essential role in intracellular K
336
homeostasis by controlling the K vacuolar release (Hamamoto and Uozumi, 2014).
337
Calcium is usually present at high concentrations in the cytosol of root cells, where it
338
has a pivotal role in the stabilization of cell walls, and also in the vacuole, where its
339
content is regulated by a larger number of Ca transporters (Pittman, 2011). The
340
cytosolic concentration of Ca is around 100 nM while the vacuolar concentration is in
341
the mM range. Hence, vacuolar compartmentation of Ca is an energy-dependent process
342
mediated by two classes of Ca vacuolar transporters (Martinoia et al., 2012). The first
343
class includes the P-type Ca-ATPases and includes two Ca transporters, ACA4 and
344
ACA11, localized in the vacuolar membrane. ACA4 occurs mainly on small vacuoles,
345
whereas ACA11 is localized in the large vacuoles of root cells (Lee et al., 2007). The
346
second class of vacuolar Ca transporters includes the Ca2+/H+ exchangers (CAXs). Six
347
CAX transporters have been identified in Arabidopsis thaliana that are split into two
348
phylogenetic groups that seems to reflect their functional variation (Manohar et al.,
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2011). The Type-1B group, which includes CAX2, CAX5 and CAX6, seem to have a
350
broader substrate specificity confirmed by its ability to transport several divalent cations
351
(e.g., Mn and Cd) besides Ca (Edmond et al., 2009). The Type 1A group, including the
352
CAX1, CAX3 and CAX4 transporters, appears to have a higher specificity for Ca
353
transport (Cheng et al., 2005). Arabidopsis cax1 knockout mutants do not suffer
354
significant changes in their Ca content, however, cax1cax3 double mutants exhibited
355
reduced Ca concentration in their vacuoles compared to wild-type plants (Conn et al.,
356
2011b, Connorton et al., 2012).
357
Plants growing on Mg-rich soils use this divalent cation as an osmotic regulator. The
358
MGTs (Mg transporters) are the major transporters of this cation described in plants. In
359
Arabidopsis thaliana, ten members with Mg2+/H+ antiport activity are identified
360
(Martinoia et al., 2012). Two of them, AtMGT2/AtMRS2-1 and AtMGT3/AtMRS2-5,
361
are localized in the vacuolar membrane. Plant lines overexpressing these transporters
362
have shown Mg accumulation (Conn et al., 2011a). Arabidopsis Mg2+/H+ exchanger
363
(AtMHX) is another putative vacuolar Mg2+/H+ antiporter that seems to mediate the Mg
364
influx across the tonoplast (Shaul et al., 1999). The same mechanism was also observed
365
in the hyperaccumulator Arabidopsis halleri expressing an AtMHX homolog (Elbaz et
366
al., 2006).
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368
Trace essential elements
369
Plants require certain micronutrients for proper growth and development. For example,
370
Fe is involved in numerous redox reactions, Cu has a crucial role in electron transfer
371
between photosystem I and II, Mn is required as a cofactor for a variety of enzymes,
372
such as the Mn-dependent superoxide dismutase (MnSOD), and Zn acts as a catalytic or
373
structural co-factor in a large number of enzymes and regulatory proteins (Hänsch and
16 Page 16 of 78
Mendel, 2009). Micronutrients deficiency is a more common reality that toxicity.
375
Hence, plants usually accumulate extra nutrients to sustain their growth in times of
376
need. To accomplish that, micronutrients are translocated to seeds where they can
377
support the growth of new plantlet. Therefore, most of these micronutrients are not
378
retained in plant roots, so this topic is less relevant to discuss. Only a brief overview
379
will be made here.
380
The bulk of Fe is found in leaves chloroplasts, where this element is engaged in several
381
biochemical reactions (Hänsch and Mendel, 2009). A certain amount of this
382
micronutrient can also be stored in the leaves vacuoles or mitochondria (Martinoia et
383
al., 2012). Iron can also be stored in seeds vacuoles (Kim et al., 2006b). Its storage in
384
root cells is not very common.
385
Non-tolerant plant species, such as A. thaliana, can use chelation as a mechanism to
386
avoid Cu-induced damage in the root cells. In a Cu-rich environment, MTs and PCs are
387
usually up-regulated in response to Cu stress (Guo et al., 2008, Pal and Rai, 2010). Four
388
Arabidopsis MTs (AtMT1a, AtMT2a, AtMT2b, and AtMT3) have been implicated in
389
the Cu accumulation and tolerance in roots under high Cu levels (Guo et al., 2008). In
390
addition to the MTs tolerance mechanism, Cu can be sequestered in the vacuole. A
391
member of the HMA (Heavy Metal Associated) family, AtHMA5, mediates the
392
vacuolar Cu influx in Arabidopsis root cells (Andres-Colas et al., 2006). In A. thaliana,
393
one of the six COPTs identified is targeted to the tonoplast. An Arabidopsis copt5
394
knockout mutant exhibits severe defects in vegetative growth and root elongation,
395
accumulating more Cu in the roots and translocating less Cu to the shoots. These results
396
support the fact that AtCOPT5 would act as a vacuolar Cu exporter (Garcia-Molina et
397
al., 2011, Klaumann et al., 2011).
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A large pool of Mn (around 80%) is usually observed in leaf vacuoles. Moreover, a
399
significant accumulation of Mn is also present in leaf chloroplasts where this element is
400
required as the catalytic center for light-induced water oxidation in photosystem II
401
(Pittman, 2005). The previously mentioned AtCAX2 and AtCAX4 transporters that are
402
engaged in the transport of Ca have also the ability to transport Mn into the vacuole
403
(Edmond et al., 2009, Connorton et al., 2012). AtMTP11, which belongs to the cation
404
diffusion facilitator (CDF) family, is implicated in the pre-vacuolar compartmentation
405
of Mn in root cells (Delhaize et al., 2007). In Arabidopsis, the export of Mn from
406
vacuoles can be performed by AtNRAMP3 and AtNRAMP4 which were shown to have
407
a crucial role in the Mn homeostasis within plant tissues (Lanquar et al., 2010). The P-
408
type ATPase AtECA3, mainly expressed in root tissues, participates in the influx of Mn
409
into the Golgi complex (Mills et al., 2008).
410
After uptake into the root symplast, Ni detoxification at the cellular level is achieved by
411
transport of Ni, as a free ion or chelated with appropriate ligands, to the storage sites
412
such as the vacuole and cell wall (Bhatia et al., 2005). A recent study showed that the
413
main mechanism that confers Ni tolerance seems to be vacuolar Ni sequestration (Saito
414
et al., 2010). A member of the IREG/FPN (Iron-Regulated/Ferroportin) family,
415
AtIREG2/FPN2, has been identified in the tonoplast of root cells and mediates vacuolar
416
Ni loading (Schaaf et al., 2006).
417
Once in the root symplast, Zn can be immobilized in the root through transport into
418
vacuoles. Evidence suggests that the speciation of Zn in the root symplast has a direct
419
influence in the extent of Zn immobilization in root vacuoles (Sinclair and Kramer,
420
2012). The vacuolar membrane transporter AtMTP1 has an essential role in
421
detoxification of excessive Zn. When grown in a Zn-rich medium, the mutant line of
422
Arabidopsis that lacks MTP1 was not able to accumulate Zn in vacuoles, unlike wild-
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type roots (Kawachi et al., 2009). The tonoplast-localized AtMTP3 also contributes to
424
basic cellular Zn tolerance and controls Zn partitioning. Overexpression of MTP3
425
increases Zn accumulation in roots of A. thaliana (Arrivault et al., 2006). The P1B-type
426
ATPase subfamily HMA plays a key role in the process of metal allocation or
427
detoxification in plants. At least eight (HMA1-HMA8) members are identified in
428
Arabidopsis thaliana. AtHMA1 to AtHMA4 belong to the group implicated in divalent
429
cation transport whereas AtHMA5 to AtHMA8 act on monovalent Cu ion transport
430
(Zorrig et al., 2011). From the four HMAs involved in divalent cations transport, one of
431
them, AtHMA3, seems to play a role in the detoxification of Zn by participating in its
432
vacuolar sequestration in root cells (Morel et al., 2009). It has been speculated that ZIF1
433
(for Zinc-Induced Facilitator1) is involved in a mechanism of vacuolar Zn sequestration.
434
Specifically, AtZIF1 has been implicated in the transport of Zn complexes (mainly with
435
NA) into the vacuole, since overexpression of ZIF1 leads to strongly enhanced retention
436
of Zn in root vacuoles (Haydon et al., 2012). A member of the ZIP family, AtZIP1, may
437
contribute to the remobilization of Zn from the vacuole to the cytoplasm in Arabidopsis
438
roots (Milner et al., 2013). Moreover, the tonoplast-localized NRAMP4 also seems to
439
mediate the vacuolar Zn export in root cells (Oomen et al., 2009).
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423
441
Beneficial elements
442
Sodium is a ubiquitous cation present in all soils. Soils can contain high concentrations
443
of Na, which can be readily absorbed by most plants. Since high Na concentrations
444
within the cytosol can be toxic, vacuolar Na storage is essential for plants to survive
445
under high salinity conditions (Martinoia et al., 2012, Benito et al., 2014). As already
446
described, the NHX family is a Na+/H+ antiporter. Several member of the NHX family
447
(NHX1-4) are targeted in the vacuole and are known to be involved in salt tolerance.
19 Page 19 of 78
Furthermore, the endosomal-localized NHX5 and NHX6 also seem to be engaged in
449
this process, suggesting a complex interactive coordination between different cell
450
organelles in response to salinity (Krebs et al., 2010, Bassil et al., 2011).
451
As regard to Co, few studies have addressed its compartmentation. In fact, the only
452
evidence about vacuolar compartmentation of Co comes from the study of Morrissey et
453
al. (2009) who found that AtIREG2/FPN2, besides being involved in the vacuolar
454
sequestration of Ni, also seems to mediate Co transport across the tonoplast in root
455
cells.
us
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448
an
456
Translocation
458
Translocation in plants involves a series of steps, as follows: (1) radial transport across
459
the root tissues, which include symplastic transport to pass through the Casparian strip
460
(a waxy coating that prevents solutes from entering the root xylem directly from the soil
461
solution or root apoplast); (2) xylem loading, transport and unloading; (3) xylem-to-
462
phloem transfer; (4) phloem loading, transport and unloading; (5) symplastic movement
463
toward the site of demand; and (6) retranslocation from source or senescing tissue
464
(Kobayashi and Nishizawa, 2012).
465
Although some transporters are highly specialized in the movement of a certain cation,
466
others show low substrate specificity and can translocate different elements (Zorrig,
467
Abdelly, 2011). Some cations are chelated by organic acids, which are involved in metal
468
absorption by plant roots, translocation in the xylem and storage in the leaf cells
469
vacuoles (Karley and White, 2009, Larbi et al., 2010). Cations can also be bound by NA
470
and mobilized by the YSL family transporters (Aoyama et al., 2009, Ishimaru et al.,
471
2010, Harris et al., 2012). It should be emphasized that cation translocation is a complex
472
process involving multiple networks. Specialized proteins can transport elements with
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473
similar characteristics (e.g., oxidative state), inducing competition that can results in
474
deficiency, toxicity and/or accumulation in the above-ground plant tissues (Puig and
475
Penarrubia, 2009).
ip t
476
Macroelements
478
A large fraction of the K taken up by plant roots is translocated to the shoot. Potassium
479
is transported symplastically to the xylem, where the Shaker-like channel AtSKOR,
480
expressed in the root pericycle and stelar parenchyma of Arabidopsis, is thought to be
481
involved in K loading of the xylem (Johansson et al., 2006). The loading of K within the
482
shoot is mainly determined by transpirational water flows and the shoot apoplastic K
483
concentration (Ahmad and Maathuis, 2014). In addition to the transport of K into
484
above-ground tissues, a large shoot-to-root K flux through the phloem is maintained. In
485
Arabidopsis, the redistribution of K from leaves to phloem-fed tissues seems to be
486
mediated by the Shaker-like channel AtAKT2/AKT3, which is mainly expressed in the
487
phloem and xylem parenchyma and has been indicated to be involved in phloem loading
488
and unloading (Gajdanowicz et al., 2011).
489
Calcium is relatively immobile within plant tissues and thus, it is likely that most of the
490
Ca is sequestered in the root cells vacuoles (Maathuis, 2009). Until now, no transporters
491
have been identified to be responsible for Ca xylem loading, although it is possible for
492
Ca to be symplastically transported across the root to the xylem (Karley and White,
493
2009). Evidence suggests that Ca reaches the xylem apoplastically through regions of
494
the root where the Casparian strips are absent or via the cytosol of unsuberised
495
endodermal cells where Casparian strips are present (Baxter et al., 2009). This
496
assumptions support the observation that Ca mobilization to the shoot occurs mainly
497
from the root apex and/or regions of lateral root initiation, where the Casparian strip is
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disrupted (Shaul, 2002, White et al., 2002). Once in the vascular system, Ca mobility is
499
also low. Calcium is transported in the xylem as free ions (Ca2+) or complexed with
500
organic acids (Karley and White, 2009). Additionally, it seems that the rhizosphere Ca
501
concentration has an important effect on the Ca concentration in the xylem sap and
502
transpirational water flows governs Ca delivery within the shoot (White et al., 2002).
503
Magnesium is loaded into the xylem sap, where it is present as free ions (Mg2+) or
504
complexed with organic acids (Karley and White, 2009). To date, little information is
505
available regarding membrane transporters engaged in the Mg xylem loading. Recently,
506
it was suggested that members of the MRS2/MGT family could be involved in the Mg
507
transport system (Tanoi et al., 2011), but the exact transporter has not been identified
508
yet. As for Ca, evidence suggests that, besides the symplastic pathway, Mg can be
509
loaded into the xylem vessels apoplastically and it seems that this pathways is the
510
preferred for xylem Mg loading (Shaul, 2002).
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498
te
511
Trace essential elements
513
Within plant tissues, Fe has low solubility and high reactivity thus its movement along
514
the plant tissues must be associated with appropriate chelating compounds and proper
515
control of redox states (Kobayashi and Nishizawa, 2012). Several lines of evidence have
516
indicated that the main Fe chelators are citrate (Rellan-Alvarez et al., 2010, Harris et al.,
517
2012), NA (Ishimaru et al., 2010) and MAs (Aoyama et al., 2009, Kakei et al., 2012).
518
Citrate has been proved to play a key role in the Fe chelation and mobilization in xylem
519
sap. To facilitate the efflux of citrate into the xylem, Arabidopsis relies on the FRD3
520
(Ferric Reductase Defective 3) transporter which is expressed in the root pericycle cells
521
(Figure 5) (Durrett et al., 2007). In rice, an AtFDR3 orthologue, OsFRDL1, is also
522
expressed in root pericycle cells and encodes a citrate effluxer required to load citrate
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into the xylem sap (Yokosho et al., 2009). OsPEZ1 and OsPEZ2 are also responsible for
524
xylem loading of phenolics enhancing the remobilization of Fe along the plant body
525
(Bashir et al., 2011a, Ishimaru et al., 2011). Although AtFRD3, OsFRDL1, OsPEZ1 and
526
OsPEZ2 can efflux the Fe-chelating molecules, Fe must also be transported into the
527
xylem. To date, the plasma membrane-localized IREG1/FPN1 present in Arabidopsis
528
stele, is the only Fe effluxer identified to mediate the vascular loading of Fe (Morrissey
529
et al., 2009). Among the influx transporters, members of the YS and YSL transporters
530
are widely involved in Fe translocation. In barley and maize, the YS1 transporter is able
531
to mobilize Fe-PS complexes (Murata et al., 2008, Ueno et al., 2009). Among the 18
532
YSL transporters present in rice, OsYSL2, OsYSL15, OsYSL16, OsYSL18 were
533
identified as Fe-PS transporters involved in the Fe translocation. More specifically,
534
OsYSL2 is proved to be a Fe-NA transporter, whereas OsYSL15, OsYSL16, OsYSL18
535
translocate Fe-deoxymugineic acid (DMA) complexes (Aoyama et al., 2009, Inoue et
536
al., 2009, Ishimaru et al., 2010, Kakei et al., 2012). The efflux of NA in rice was found
537
to be facilitated by the ENA1 (efflux transporter of nicotianamine 1) and ENA2, and the
538
efflux of MA is achieved by TOM1 (Nozoye et al., 2011). Members of the YSL family
539
are also present in non-graminaceous plants that do not synthesize MAs. It is widely
540
assumed that non-graminaceous plants rely on the YSL transporters to translocate Fe-
541
NA complexes. NA is a precursor and structural analogue of MAs; it has the ability to
542
chelate Fe and other metals and is synthesized by all plants, in contrast to the
543
graminaceous-specific synthesis of MAs. The Arabidopsis YSL members AtYSL1,
544
AtYSL2 and AtYSL3 were identified to have a key role in the mobilization of Fe inside
545
the plant (Chu et al., 2010). Besides their presence in the outer root cells, some FRO
546
genes can also be expressed in root or shoot vascular cells (Jeong and Connolly, 2009),
547
suggesting that Fe reduction can occur within plant tissues with further Fe translocation
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23 Page 23 of 78
by members of the IRT and/or NRAMP families (Takahashi et al., 2011). The
549
oligopeptide transporters (OPT) seem to play an important role in the mobilization of
550
Fe. OPT3 was markedly expressed in aerial parts of both Arabidopsis thaliana and
551
Thlaspi caerulescens, including stem and leaf, suggesting that this transporter may be
552
involved in the long-distance Fe transportation (Stacey et al., 2008, Hu et al., 2012).
553
In order to be translocated, Cu must be exported from the root symplast before entering
554
the xylem. Evidence suggests that the export from the root symplast is mediated by an
555
HMA family transporter. In Arabidopsis, AtHMA5 is highly expressed in roots, flowers
556
and pollen and seems to be responsible for root Cu detoxification (Andres-Colas,
557
Sancenon, 2006). This phenotype function is the opposite of COPT, corroborating the
558
idea that AtCOPT1 and AtHMA5 transport Cu in opposite directions (Figure 5) (del
559
Pozo, Cambiazo, 2010). Cu translocation may involve chelators such as NA and several
560
amino acids (Irtelli et al., 2009, Harris et al., 2012). Moreover, Cu in the xylem sap of
561
rice seems to be bound to DMA, while in the phloem sap, Cu mainly complexes with
562
NA and histidine (Ando et al., 2013). OsYSL16 is a Cu-NA transporter required for
563
delivering Cu to developing young tissues and seeds through phloem transport (Zheng
564
et al., 2012). In Arabidopsis root, the plasma membrane-localized AtYSL1 and AtYSL3
565
were proven to be involved in the root-to-shoot Cu transport (Waters and Grusak, 2008,
566
Chu et al., 2010). In Arabidopsis, AtCOPT6 is expressed in different cell types of
567
different plant compartments, but the bulk of its expression is located in the vasculature
568
suggesting a role in Cu translocation (Jung et al., 2012). Similarly, OsCOPT6 is not
569
expressed in root but is highly expressed in leaf, stem and sheath (Yuan et al., 2011).
570
Furthermore, the oligopeptide transporter AtOPT3 and its orthologue TcOPT3 seem to
571
participate in the mobilization of Cu (Stacey et al., 2008, Hu et al., 2012).
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24 Page 24 of 78
The translocation of Mn involves chelators such as NA, amino acids and carboxylic
573
acids (Harris et al., 2012). In Arabidopsis, AtZIP2 seems to participate in Mn transport
574
into the root vasculature for translocation to the shoot (Figure 5) (Milner et al., 2013). In
575
rice, OsYSL6 is a Mn-NA transporter that is required for the detoxification of excess
576
Mn (Sasaki et al., 2011). Likewise, OsYSL2 is involved in Mn translocation in the
577
phloem. In the knockout ysl2 line, rice seeds and shoots contain lower Mn content
578
compared with the wild-type (Ishimaru et al., 2010). Moreover, the same transporter
579
(YSL2) shows the ability to transport Mn-PS complexes in barley roots (Araki et al.,
580
2011). The oligopeptide transporter AtOPT3 seems to play an important role in the
581
long-distance Mn transport (Stacey et al., 2008).
582
Nickel is translocated from roots to shoots as a free ion (Ni2+) and complexed with
583
several molecules such as histidine and citrate (Richau et al., 2009, Centofanti et al.,
584
2013). In hyperaccumulator plants, Ni was shown to be translocated also in the form of
585
Ni-NA complexes (Mari et al., 2006). The transport of NA-metal chelates, required for
586
the xylem loading and unloading, has been assigned to the YSL family. In Thlaspi
587
caerulescens, two YSL genes, TcYSL5 and TcYSL7 were found to be expressed around
588
the vasculature of the shoots and in the central cylinder of the roots, suggesting that
589
YSL transporters play a key role in the translocation of Ni-NA complexes (Gendre et
590
al., 2007).
591
Zinc is exported from root by two members of the P-type ATPase family, HMA2
592
(expressed in Arabidopsis, rice and barley) and HMA4 (Figure 5) (Mills et al., 2012,
593
Takahashi et al., 2012b). However, other transporters can also perform the export of Zn
594
to vascular tissues. In Arabidopsis, pcr2 knockout mutants accumulate Zn in roots,
595
suggesting a role of AtPCR2 in the root-to-shoot Zn translocation (Song et al., 2010).
596
Furthermore, it was proved that AtFRD3 is involved in loading Zn into xylem (Pineau
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25 Page 25 of 78
et al., 2012). Zn can also be exported from the roots in the form of Zn complexes.
598
Before being exported, Zn must be mobilized to pericycle cells, apparently by HvYSL2
599
(Araki et al., 2011). After xylem loading, Zn can be transported to above-ground tissues
600
by the YS and YSL transporters. There is some evidence that AtYSL1, AtYSL2 and
601
AtYSL3 are involved in the transport of Zn (Chu et al., 2010). Phytosiderophores such
602
as NA and DMA are important in the distribution of Zn within the plant (Suzuki et al.,
603
2008, Harris et al., 2012, Nishiyama et al., 2012). Although no Zn-PS transporters have
604
been identified yet, the presence of these complexes in the xylem and phloem sap
605
suggests that members of the YSL family may be involved in the transport of Zn
606
complexes (Rellan-Alvarez et al., 2008, Deinlein et al., 2012). Moreover, members of
607
the ZIP family are expressed in the vascular bundle, suggesting that they contribute to
608
the distribution of Zn along the plant (Bashir et al., 2012). AtOPT3 and its orthologue
609
TcOPT3 seem to participate in the long-distance transport of Zn (Stacey et al., 2008, Hu
610
et al., 2012).
te
611
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597
Beneficial elements
613
The export of Na to the xylem is assumed to be an active process. In Arabidopsis, SOS1
614
was proposed to be involved in the loading of Na into the xylem (Shi et al., 2002).
615
However, at the whole-plant level, the specific role of AtSOS1 still remains uncertain
616
(Kronzucker and Britto, 2011). At high Na levels, xylem loading with Na is thought to
617
occur passively through channels, since a higher cytosolic Na content in xylem
618
parenchyma cells and a relatively depolarized plasma membrane would promote the
619
movement of Na into the xylem sap (Apse and Blumwald, 2007). Several lines of
620
evidence exist supporting the role of AtHKT1;1 and its orthologue OsHKT1;5 in xylem
621
unloading (Sunarpi et al., 2005, Davenport et al., 2007). A demonstration of Na efflux
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26 Page 26 of 78
from shoot xylem vessels was achieved by overexpressing AtHKT1;1 in the mature root
623
stele of A. thaliana, which showed decreased Na accumulation in the shoot (Moller et
624
al., 2009).
625
For Co, little evidence is available regarding its translocation. However, an Arabidopsis
626
fpn1 knockout mutant showed low Co accumulation in shoot tissues, indicating that
627
ferroportins play an important role in Co homeostasis, more specifically in the root-to-
628
shoot Co translocation (Morrissey et al., 2009).
us
629
cr
ip t
622
Leaves compartmentation
631
The transport of inorganics into leaf cells from leaf xylem involves another membrane
632
transport step mediated by specific membrane transport proteins. Once inside the leaf
633
symplast, cations can be assimilated in the cytosol or stored in cellular organelles. The
634
general rule is that nutrients are assimilated, playing a key role in several physiological
635
mechanisms, and toxic elements are stored in places where they cause the least harm to
636
cellular processes. However, it should be noted that nutrients can also be stored for two
637
reasons: (i) they can become deficient and plant accumulates them for times of
638
necessity; (ii) they are at high concentrations and can pose a toxicity problem to plants
639
(Maathuis, 2009, Pilon-Smits et al., 2009, Pilon et al., 2009). At the tissue level, toxic
640
elements are generally accumulated in the epidermis and trichomes (Tian et al., 2009,
641
Sanchez-Pardo et al., 2012); at the cellular level, they may be accumulated in the
642
vacuoles or cell wall (Cosio et al., 2005).
643
The distribution of cations in leaf tissues is generally asymmetric (Wu and Becker,
644
2012). The accumulation of toxic cations in leaf vacuoles makes sense because vacuoles
645
do not contain a photosynthetic apparatus, which would be sensitive to toxicity
646
(Martinoia et al., 2012). Toxic cation storage occurs also in highly tolerant cells such as
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the leaf epidermis and the vein bundle sheath, as long as their import remains under
648
control. Once the cation accumulation exceeds the tolerance threshold of the plant, they
649
would be transported to mesophyll cells, which are more sensitive to toxic cations than
650
other cell types, allowing photosynthesis to be threatened (Zhao et al., 2012). At
651
present, the physiological mechanisms involved in the sequestration of cations between
652
different leaf tissues remain only partially understood. One possible mechanism is the
653
differential expression of cation transporters in the plasma membrane and/or tonoplast
654
between mesophyll and other tissues (Morel et al., 2009).
us
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655
Macroelements
657
The transport of K from the xylem into the leaf symplast is mediated by membrane
658
transporters and channels. Members of the HAK/KT/KUP family were found to be
659
highly expressed in shoot tissues. Specifically, AtKUP1, AtKUP2 and AtKUP4 were
660
expressed in Arabidopsis leaves, suggesting an involvement in K xylem unloading
661
(Wang and Wu, 2013). Moreover, Ahn et al (2004) also showed that other AtKUPs
662
were highly expressed in leaves. In particular, AtKUP6, AtKUP8 and AtKUP9 were
663
highly expressed in young leaves, whereas AtKUP10 and AtKUP12 expression was
664
evident in both older and younger leaves. Shaker channels AtKT1, AtKT2 and AtKC1
665
are also highly expressed in the plasma membrane of Arabidopsis leaves, indicating that
666
these cation channels are enrolled in the movement of K between shoot tissues
667
(Jeanguenin et al., 2011). Regarding the intracellular distribution of K in leaves, the
668
vacuolar K transporters AtNHX1 and AtNHX2, expressed in the tonoplast of
669
Arabidopsis leaf vacuoles, mediate the influx of K into this cell organelle (Figure 6)
670
(Bassil et al., 2011). To mediate the vacuolar K efflux, several members of the
671
TPK/KCOs family, such as AtTPK1, AtTPK3, AtTPK5 and AtKCO3, are expressed in
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the vacuolar membrane of leaf vacuoles and mediate this process (Voelker et al., 2010,
673
Hamamoto and Uozumi, 2014). Regarding other cell organelles, a member of the CHX
674
family, AtCHX23, was found to be expressed in the chloroplast envelope of
675
Arabidopsis leaf cells. Analysis of Arabidopsis chx23 knockout mutant leaves suggests
676
that CHX23 could mediate the co-transport of K across the chloroplasts envelope of leaf
677
cells (Song et al., 2004). Recently, a member of the K efflux antiporters (KEA) has also
678
been identified in Arabidopsis chloroplasts. AtKEA2 was found to be highly expressed
679
in leaves and mediates K transport across the plastid envelope (Aranda-Sicilia et al.,
680
2012).
681
Within the leaf, Ca follows mainly the apoplastic pathway and tends to accumulate in
682
the mesophyll cells, trichomes and epidermal cells. The cytosolic Ca concentration is
683
very sensitive to the apoplastic Ca concentrations. Thus, Ca is usually stored in leaf
684
vacuoles and/or specific cell organelles to avoid excessive apoplastic Ca accumulation
685
(White and Broadley, 2003). However, before been sequestered in the leaf vacuoles, Ca
686
must be transported across the plasma membrane of leaf cells. Although most members
687
of the NSSC are present in the root cells, several lines of evidence suggest that VI-
688
NSCCs can also be found in leaf cells (Demidchik and Maathuis, 2007). In the vacuoles
689
of leaf cells, Ca is present as the ionic species Ca2+, in soluble forms (e.g., complexed
690
with proteins and/or organic acids) and/or in insoluble forms (e.g., Ca-oxalate and Ca-
691
phytate) (Karley and White, 2009). The transport of Ca across the vacuolar membrane
692
of leaf cells is thought to be performed by members of the ACA and CAX families
693
(Figure 6). AtACA11 is targeted to the vacuolar membrane of Arabidopsis leaf cells and
694
mediates Ca transport across the tonoplast of large vacuoles (Lee et al., 2007). In the
695
CAX family, AtCAX1 is highly expressed in the vacuoles of leaf cells and has an
696
important role in Ca influx into the vacuole (Cheng et al., 2005, Conn et al., 2011b).
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AtCAX2, AtCAX5 and AtCAX6 were also expressed in Arabidopsis leaves, although in
698
a lower extent than AtCAX1 (Edmond et al., 2009).
699
Members of the MGT/MRS2 family are thought to mediate the leaf uptake of Mg
700
(Gebert et al., 2009, Karley and White, 2009). It was shown that AtMGT9 is expressed
701
in leaf cells and thus can be involved in Mg transport (Chen et al., 2009). Once in the
702
cytosol of leaf cells, Mg can be accumulated in several cell organelles. Evidence
703
suggests that mesophyll cells accumulate the highest vacuolar concentration of Mg in
704
Arabidopsis leaves. To accomplish that, plants rely on members of the MGT/MRS2
705
family, such as AtMGT2/MRS2-1 and AtMGT3/MRS2-5, which are localized in the
706
tonoplast of Arabidopsis leaf cells and mediate the Mg influx into the vacuole (Figure
707
6) (Conn et al., 2011a). AtMHX is also involved in Mg influx across the tonoplast of
708
leaf cells (Shaul et al., 1999, Elbaz et al., 2006). A member of the MGT/MRS2 family,
709
AtMRS2-11, was expressed in the chloroplast envelope and plays a key role in the Mg
710
transport into this cell organelle (Drummond et al., 2006).
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Trace essential elements
713
Fe can be transported into the leaf symplast as free ion or complexed with PS. Members
714
of the IRT family are expressed in leaf tissues of Arabidopsis suggesting their
715
involvement in the Fe acquisition (Lee and An, 2009, Lin et al., 2009, Barberon et al.,
716
2011). Regarding Fe-PS complexes, members of the YSL family seems to mediate the
717
Fe influx into the cytosol of leaf cells (Kakei et al., 2012). Once in the leaf symplast, Fe
718
must be delivered into leaf cells to participate in multiple biochemical processes. The
719
chloroplast is the largest pool of Fe in plant cells accumulating 80 to 90% of cellular Fe
720
(Kobayashi and Nishizawa, 2012). The influx of Fe into the chloroplast of Arabidopsis
721
thaliana is achieved by the PIC1 (permease in chloroplast 1) localized in the inner
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envelope membrane of chloroplasts (Figure 7) (Duy et al., 2011). AtFRO7 is localized
723
in the chloroplast envelope and looks to be critical for chloroplast Fe acquisition (Jeong
724
and Connolly, 2009). Mitochondria are other plant sites where Fe is essential. Bashir et
725
al. (2011b) identified the rice mitochondrial Fe transporter (MIT), which is essential for
726
rice growth and development. Moreover, AtFRO3 and AtFRO8 are localized in
727
mitochondria membrane, which indicates that these ferric reductase oxidases must be
728
involved in mitochondrial Fe homeostasis (Jeong and Connolly, 2009). The vacuole
729
generally functions as a metal pool to avoid toxicity. The transporters OsVIT1 and
730
OsVIT2 mediate the influx of Fe across the vacuolar membrane into the vacuole (Zhang
731
et al., 2012). Likewise, it seems that the IREG2/FPN2 present in Arabidopsis tonoplast
732
may play a role in the influx of Fe from the cytoplasm into the vacuole (Morrissey et al.,
733
2009). On the other hand, both AtNRAMP3 and AtNRAMP4 export Fe from vacuoles
734
(Lanquar et al., 2005). In the Golgi complex, the MATE (multidrug and toxic
735
compound extrusion) transporter BCD1 has a role in sustaining Fe homoeostasis by
736
reallocating excess Fe released from stress-induced cellular damage in Arabidopsis (Seo
737
et al., 2012).
738
In order to Cu reach the leaf symplast, plasma membrane proteins must exist to mediate
739
Cu xylem unloading and Cu acquisition. It seems that members of the COPT family are
740
responsible for the influx of monovalent Cu (Sancenon et al., 2004, Andres-Colas et al.,
741
2006). However, since Cu is present mainly in the divalent form in the vascular tissues,
742
reduction must occur before it can enter the cell organelles. AtFRO7 (expressed in the
743
chloroplasts) and AtFRO3/8 (expressed in the mitochondria) play a central role in Fe
744
reduction, and it is hypothesized that they also participate in Cu reduction (Jeong and
745
Connolly, 2009). Moreover, Cu can also be transported across the plasma membrane of
746
leaf cells complexed with PS. Members of the YSL family were proved to be engaged
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in the Cu translocation and delivery to shoot tissues (Irtelli et al., 2009, Zheng et al.,
748
2012, Ando et al., 2013). Once in the cytosol of leaf cells, Cu is distributed with the aid
749
of plasma membrane-localized transporters among different cell organelles. AtHMA6
750
(or PAA1), localized in the inner chloroplast envelope, is responsible for the delivery of
751
Cu to chloroplasts (Figure 7) (Catty et al., 2011). AtHMA8 (PAA2), closely related to
752
AtHMA6 (PAA1), is expressed in the thylakoid membrane and supplies Cu to
753
plastocyanin (Tapken et al., 2012). AtHMA1 and HvHMA1, present in the chloroplast
754
envelope, are broad-specificity exporters of metals from chloroplasts and may play a
755
specialized role in Cu mobilization (Mikkelsen et al., 2012). AtHMA7 (RAN1), the first
756
functionally characterized heavy metal ATPase, is responsible for the biogenesis of
757
ethylene receptors by supplying Cu at the endoplasmic reticulum and also for Cu
758
homeostasis in seedling development (Binder et al., 2010). The rice mitochondrial Fe
759
transporter (MIT) also appears to regulate the influx of Cu, although more studies are
760
needed to confirm this (Bashir et al., 2011b). The tonoplast-localized AtCOPT5 is
761
important for Cu export from the vacuole and is involved in the remobilization of Cu
762
ions (Garcia-Molina et al., 2011, Klaumann et al., 2011).
763
Manganese reaches the leaf symplast by crossing the plasma membrane of leaf cells. To
764
do that, members of the IRT family are thought to be expressed in the shoot tissues,
765
mediating the transport of Mn ions into the cytosol of leaf cells (Pedas et al., 2008,
766
Barberon et al., 2011). After influx into the leaf symplast, Mn must be distributed into
767
various cell compartments. In Arabidopsis, AtECA1 is localized at the endoplasmic
768
reticulum (Li et al., 2008). The cation/H+ exchanger (CAX) transporters AtCAX2 and
769
AtCAX4, which were originally identified as Ca2+ transporters, also have the ability to
770
transport Mn into the vacuole (Figure 7) (Connorton et al., 2012). Likewise, vacuolar
771
Mn2+/H+ antiporter activity in the Arabidopsis cax2 knockout mutant is significantly
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reduced, although it is not completely absent, compared with wild-type, suggesting the
773
presence of additional vacuolar transporters (e.g., AtCAX5) that contribute to Mn
774
transport (Edmond et al., 2009). AtMTP11, which belongs to the cation diffusion
775
facilitator (CDF) family, is implicated in the pre-vacuolar compartmentation of Mn as
776
well as in Mn homeostasis mechanisms (Delhaize et al., 2007). Other MTPs also
777
contribute to Mn transport (Gustin et al., 2011). Furthermore, it seems that the
778
tonoplast-localized transporters OsVIT1 and OsVIT2 participate in Mn influx to the
779
vacuole (Zhang et al., 2012). Regarding vacuolar export, AtNRAMP3 and AtNRAMP4
780
are expressed in the tonoplast and can transport Mn and other metals (Lanquar et al.,
781
2010). AtZIP1 probably plays a role in Mn vacuolar efflux, based on the increased
782
sensitivity to low Mn and increased accumulation of Mn in roots in the zip1 knockout
783
line (Milner et al., 2013). The rice Fe transporter MIT appears to play a role in Mn
784
transport into mitochondria (Bashir et al., 2011b). The mechanisms of Mn transport in
785
the chloroplast remain unknown, despite the fact that Mn has a critical role in
786
photosynthesis and the chloroplast is the one of the major sinks for Mn (Yao et al.,
787
2012, Millaleo et al., 2013).
788
Zinc enters the leaf symplast with the aid of members of the IRT and ZIP families
789
(Sinclair and Kramer, 2012). After reaching the leaf cytosol, the delivery of Zn to plant
790
organelles is also mediated by specific transporters (Figure 7). The vacuolar membrane
791
transporters AtMTP1 and AtMTP3 are expressed in the vacuoles of leaf cells and play a
792
key role in detoxification of excessive Zn by accumulating Zn in vacuoles (Arrivault et
793
al., 2006, Gustin et al., 2009). Likewise, AtHMA3 is also involved in vacuolar Zn
794
sequestration (Morel et al., 2009). In Arabidopsis, AtZIF1 also seems to be expressed in
795
the tonoplast of leaf cells. Therefore, an active role is expected from this transporter
796
regarding the vacuolar Zn sequestration (Haydon et al., 2012). The tonoplast-localized
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OsVIT1 and OsVIT2 seem to transport Zn into the vacuoles of plant cells (Zhang et al.,
798
2012). In the chloroplasts of both Arabidopsis and barley, HMA1 contributes to Zn
799
detoxification by exporting Zn from the plastids to the cytoplasm (Kim et al., 2009,
800
Mikkelsen et al., 2012). It has been proposed that the Golgi-localized transporter
801
AtECA3 may have a role in the mobilization of Zn from the cytoplasm to the Golgi
802
complex (Mills et al., 2008).
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797
803
Beneficial elements
805
Regarding the beneficial cations Na and Co, little information is available about the
806
processes regulating their movement into the leaf symplast as well as their distribution
807
in the cell organelles (Pilon-Smits et al., 2009). Although their essentiality is not yet
808
proved, evidence suggests that similar membrane transporters should mediate their
809
mobilization within the plant compartments (Hänsch and Mendel, 2009, Kronzucker
810
and Britto, 2011).
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Biofortification
813
Plant foods refer to plants or parts of a plant that are eaten as a food. In this category are
814
included vegetables, fruits, grains and legumes. They constitute one of the most
815
important nutrient sources in human diet since the beginning of civilization. Humans
816
need several essential minerals in order to allow the proper functioning of the body,
817
regarding growth, development and metabolism. Recommended dietary allowances
818
(RDAs), defined as “the levels of intake of essential nutrients that, on the basis of
819
scientific knowledge, are judged by the Food and Nutrition Board to be adequate to
820
meet the known nutrient needs of practically all healthy persons” are established and
821
vary between a few μg/day and 1 g/day. If intakes are low in a certain period of time,
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deficiency signs may develop. On the other hand, high intakes can result in toxicity
823
(FNB/IOM, 2006).
824
To grow and develop, plants must take up water and essential mineral nutrients from the
825
soil. Once taken up by plant roots, nutrients can be transported and accumulate in other
826
parts of the plant. The final composition of the edible parts of plants is influenced and
827
controlled by several factors, including soil fertility, plant genetics and the environment
828
in which they grow (Karley and White, 2009, Maathuis, 2009, Puig and Penarrubia,
829
2009). Although it is possible in some cases to enhance the nutritional quality of plant
830
foods through agronomic practices and plant breeding, the soil-to-plant and plant-to-
831
human transfer of mineral nutrients is a very complex process (Hirschi, 2009). First of
832
all, soils differ greatly in their mineral composition and nutrients phytoavailability
833
depends on several factors (Palm et al., 2007, Pinto et al., 2014b). Secondly, plants
834
possess physiological mechanisms to regulate the amounts of nutrients taken up from
835
the soil. Hence, one can expect that attempts to alter the overall mineral composition of
836
plant foods will be very complex (Maathuis, 2009, Pilon et al., 2009, Pinto et al.,
837
2014a). Thirdly, it has been recognized that the concentration of a nutrient in a food is
838
not necessarily a reliable indicator of the value of that food as a source of the nutrient in
839
question. This leads us to the concept of nutrient bioavailability, defined as the amount
840
of a nutrient in ingested food that is available for utilization in metabolic processes. In
841
the case of mineral nutrients, bioavailability is determined mainly by the efficiency of
842
minerals absorption from the human gut (Hirschi, 2009). Therefore, the development of
843
plant foods biofortification must rely on a holistic approach focusing on three concepts:
844
(1) nutrients phytoavailability, (2) nutrients accumulation in edible tissues and (3)
845
nutrients bioavailability. In this review, only the two first steps (nutrients
846
phytoavailability and nutrients accumulation in edible tissues) will be discussed. For
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847
information regarding nutrients bioavailability, the reader should refer to the work of
848
Hirschi (2009).
849
Nutrients phytoavailability
851
Five areas have been identified that are related to the uptake efficiency of cations by
852
plants in which breeders can act. These include: (1) root morphology, (2) root hairs
853
formation, (3) root exudates, (4) ability to release cations from non-exchangeable pools,
854
and (5) kinetics of cation uptake. Thus, a successful biofortification strategy must
855
understand three important aspects: (1) the spatial distribution of elements within soil
856
profiles, (2) in which form elements are commonly found in the soil, and (3) how plant
857
roots acquire these elements. Gathering this knowledge is of utmost importance and will
858
enable researchers to concentrate efforts in the most limiting steps of nutrient
859
acquisition.
860
Most cations in soil are incorporated in the crystal lattice structure of minerals and thus
861
not readily available for plant uptake. The availability of these elements will be greatly
862
influenced by the soil type and its physico-chemical properties. To simplify, cations in
863
the soil can be classified in four groups depending on their phytoavailability: (1) water-
864
soluble; (2) exchangeable; (3) non-exchangeable and (4) structurally bound. Water-
865
soluble cations are directly available for both plants and microorganisms and are highly
866
mobile within soil profiles. Exchangeable cations are those weakly adsorbed elements
867
retained on the solid surface by relatively weak electrostatic interaction as well as
868
cations that can be released by ion-exchange processes (Fedotov et al., 2012). Both
869
water-soluble and exchangeable pools are usually considered to be easily
870
phytoavailable. However, the size of both pools is negligible, accounting for as much as
871
0.1-0.2% and 1–2% of the total content of these elements in soil, respectively. Non-
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exchangeable and structurally bound pools are considered to be slowly or even non-
873
available sources of elements for plants. These pools can only contribute to the supply
874
of nutrients to plants in the long term. (Zörb et al., 2014). As already described, the
875
amount of cations that are phytoavailable and non-phytoavailable in the soil vary greatly
876
among soil types. However, in all soils a dynamic equilibrium exist between the
877
different pools, which is influenced by a number of soil physico-chemical properties as
878
well as plant-soil interactions.
879
The main soil physico-chemical properties that control the processes of sorption and
880
desorption of elements in the soil are: (1) pH, (2) cation exchange capacity (CEC), (3)
881
particle size distribution, mainly the fine granulometric fraction (< 0.02 mm), (4)
882
organic matter (OM) and (5) oxides and hydroxides, mainly of Al, Fe and Mn (Kabata-
883
Pendias, 2004). Typically, the most mobile pool of elements occur at lower pH. CEC
884
reflects the dynamic equilibrium of cationic species in the soil solution, thus influencing
885
the phytoavailability of elements. When soil pH becomes acid, the phytoavailability of
886
elements usually increases due to the replacement of cations on soil binding sites by H+
887
ions. Clay minerals and oxides usually possess high active surface areas, which confer
888
them the ability to strongly bind cations. OM increases the CEC of soils because OM
889
possesses multiple binding sites for cations, which results in a great sorption capacity
890
(Pinto et al., 2014a).
891
With respect to plant-soil interactions, the rhizosphere processes such as root exudation,
892
nutrient mobilization and rhizosphere respiration, are known to greatly influence the
893
phytoavailability of elements (Hinsinger et al., 2009). Root exudation includes the
894
secretion of ions, free oxygen, water, enzymes, mucilage, and a wide variety of primary
895
and secondary metabolites (Badri and Vivanco, 2009). These exudates can increase or
896
decrease the availability of nutrients by altering soil physico-chemical properties and
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soil biological processes. Typically, these root-induced changes will alter the solubility
898
and speciation of elements in the rhizosphere in a way that nutrients are more easily
899
available for plant uptake (Badri and Vivanco, 2009, Kobayashi and Nishizawa, 2012).
900
It is now clear that one of the ways to develop more efficient plant foods, especially in
901
terms of the nutrient density, is by improving the root exudation process. This could
902
shift the equilibrium towards the enhance contribution of non-exchangeable and
903
structurally bound pools to the supply of nutrients that are many times not available for
904
plant uptake and also promote root growth as well as the formation of root hairs.
905
Breeders can start looking at the genes encoding the plasma membrane H+-ATPase
906
(HA), which are responsible for the efflux of protons that promote the acidification of
907
the rhizosphere. For instance, the breeding for an increased H+ efflux from the root in
908
order to promote the acidification of the surrounding soil may contribute to increase the
909
availability of virtually any element required for plant growth since, as above
910
mentioned, almost all elements are more mobile and thus phytoavailable at low soil pH.
911
Furthermore, H+ excretion is essential in maintaining or promoting primary root
912
elongation and root hair development (Palmgren, 2001a). Some attempts have been
913
already made in this area. In Arabidopsis, the protein kinase 5 (PKS5) and the
914
chaperone DNAJ homolog 3 (J3) have been identified to play important roles in the
915
efflux of H+ by regulating the interaction between the plasma membrane HA and 14-3-3
916
proteins (Yang et al., 2010). Furthermore, PIN2 (an auxin efflux transporter) was also
917
identified to be involved in the modulation of H+ secretion in the root tip, maintaining
918
primary root elongation (Xu et al., 2012a). More recently, it has been demonstrated the
919
involvement of 14-3-3 proteins in the regulation of H+ efflux and primary root
920
elongation. Gene specificity and functional redundancy was observed among 14-3-3
921
proteins in primary root elongation under both control and abiotic stress conditions (van
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Kleeff et al., 2014). Furthermore, TFT4 and TFT7 (tomato 14-3-3 proteins), separately
923
introduced into Arabidopsis and overexpressed, showed enhanced plasma membrane
924
HA activity, H+ efflux and also maintains elongation of the primary root (Xu et al.,
925
2012b, Xu et al., 2013a). It has been also shown that abcissic acid accumulation
926
modulates auxin transport in the root tip, which further enhances H+ secretion (Xu et al.,
927
2013b).
928
Another way to improve the nutrient density of plant foods is to manipulate the root
929
system architecture towards a distribution of roots in the soil that optimizes nutrients
930
uptake. It is important that plants have a large surface area of contact between roots and
931
soil. This allows them to have more “options” to get the water and nutrients they need.
932
The growth of the main and lateral roots is supported by cells produced in the root
933
apical meristem. In this process, phytohormones play a key role. Auxin and cytokinin
934
are assumed to be the two most important regulators of root development (Jones and
935
Ljung, 2012). The presence of auxin concentration gradients with a maximum centered
936
at the quiescent center were revealed to be essential for the establishment and
937
maintenance of the stem cell niche and for transit amplifying cell divisions (Petersson et
938
al., 2009). Cytokinin acts as an antagonist of auxin. In fact, the balance between root
939
division and differentiation appears to be primarily maintained by this antagonistic
940
relationship, with cytokinin limiting the size of the root apical meristem and the rate of
941
root growth (Zhang et al., 2011). Lateral roots are formed from pericycle cells
942
adjacently located to the xylem poles. Several genes seem to be involved in this process
943
(Moreno-Risueno et al., 2010). The identification of candidate genes will ultimately
944
allow breeders to manipulate the root system architecture to facilitate the search for
945
water and nutrients by plants. Efficient genotypes could be develop to have a relatively
946
larger proportion of thin roots as well as high number of root hairs in the whole root
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39 Page 39 of 78
system. This is thought to enhance the phytoavailability and uptake of nutrients required
948
for plant growth and development.
949
For graminaceous plants, the overexpression of genes responsible for the secretion of
950
PS will have a great impact in the solubilization of trace elements such as Fe and Zn
951
that are usually low on plant foods (Nozoye et al., 2011). Candidate genes encoding
952
ENA1 and ENA2 could be manipulated to enhance the efflux of NA in graminaceous
953
plants such as rice. Similarly, plants overexpressing TOM1 will increase the efflux of
954
MA, which could result in the increase solubilization of trace elements for further
955
uptake by plant roots.
956
Despite these improvements, it may be also necessary to manipulate the plant uptake
957
system. If increased uptake of nutrients by roots is an important goal for breeding of
958
efficient plant foods, researchers may need to take a look on the role of each nutrient in
959
plant’s metabolism. For instance, the dominant role of K is the osmoregulation. Its role
960
in turgor provision and water homeostasis is obvious in processes such as pressure-
961
driven solute transport in the xylem and phloem and high levels of vacuolar K
962
accumulation (Wang and Wu, 2013). Thus, the breeding for an increased activity of K
963
transporters/channels (e.g., AtKT/KUP or AKT1) may contribute to an increase in cell
964
turgor, leading to cell elongation during the growth of root hairs, conferring advantages
965
to plant foods growth under limiting conditions (Nieves-Cordones et al., 2014).
966
However, care should be taken to properly assess the influence of overexpressing lines
967
in maintaining the charge balance within the plant cells, which is crucial for plant
968
development. To cope with an increase in cation uptake, plants have also to increase the
969
uptake of anions like chloride, nitrate, phosphate or sulphate that also act as osmolytes.
970
This will maintain the homeostasis within plant cells. Overall, when breeding is a goal
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971
for enhance nutrient density of plant foods, ionomic studies at the whole plant level
972
should be carried out to assess the elemental distribution within plant cells.
973
Nutrients accumulation in edible tissues
975
The successful allocation of nutrients in the edible tissues of a certain plant greatly
976
depends on the plant tissue that is considered “edible”. For instance, the edible part of
977
carrots and potatoes are the roots. Therefore, in this case, attempts should be made to
978
enhance the accumulation of essential nutrients in the root of these plants. By contrast,
979
the edible tissues of leafy vegetables (e.g., lettuce or spinach) are the stems and leaves.
980
Thus, nutrients translocation must occur to move nutrients to above-ground tissues.
981
Finally, in other cases, seeds are the edible tissues of certain plants (e.g., rice) and
982
nutrients loading into seeds need to occur.
983
Two areas have been identified that are related to the utilization efficiency of elements
984
by plants in which breeders can act. These include: (1) cation translocation, and (2)
985
cation sequestration/storage. Some attempts have already been made in the two above-
986
mentioned areas. To improve cation translocation, a P1B-ATPase (AtHMA4) involved in
987
the transport of Zn in Arabidopsis was expressed in tobacco, showing enhanced Zn
988
translocation to the shoot (Siemianowski et al., 2011). Furthermore, the ectopic
989
expression of the same export protein (AtHMA4) in tobacco also resulted in decreased
990
Cd uptake/accumulation in roots and shoots (Siemianowski et al., 2014). In addition to
991
the main aim of enhancing the nutrient density of plant foods, these findings will also
992
allow breeders to manipulate plants in order to reduce the toxic exposure of humans via
993
intake of plant foods-derived Cd. Similar results have been obtained for other export
994
proteins involved in the translocation of other cations to above-ground tissues
995
(Maathuis, 2009, Pilon-Smits et al., 2009, Pilon et al., 2009, Kobayashi and Nishizawa,
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41 Page 41 of 78
2012, Mills et al., 2012, , Takahashi et al., 2012a, Ahmad and Maathuis, 2014). To
997
improve cation sequestration/storage in the organelles of edible plant tissues, breeders
998
may try to manipulate vacuolar transporters. It is now widely accepted that increasing
999
the capacity of edible tissues to sequester nutrients can promote their accumulation in
1000
these tissues. Thus, promoting the activity of vacuolar influx transporters, such as VIT,
1001
HMA, MTP and/or IREG2/FPN2, in above-ground edible tissues is seen as a promising
1002
way of food biofortification. Some research have already been made in this field. For
1003
instance, transgenic carrot and potato overexpressing the Arabidopsis vacuolar H+/Ca2+
1004
transporter AtCAX1 showed increased Ca content compared to the wild-type (Park et
1005
al., 2004, Park et al., 2005). In a similar way, potato tubers expressing the A. thaliana
1006
vacuolar transporter AtCAX2B [a variant of N-terminus truncated form of CAX2
1007
(sCAX2)] accumulate two times more Ca than wild-type (Kim et al., 2006a). Similar
1008
attempts have also been made to enhance the nutrients density of leaves and seeds. For
1009
instance, it has been shown that overexpression FRO genes results in an increase
1010
accumulation of Fe in plant leaves (Jeong and Connolly, 2009). Additionally, two
1011
important rice genes, OsVIT1 and OsVIT2, have been recently identified and
1012
characterized. It has been shown that functional disruption of both genes leads to
1013
increased Fe/Zn accumulation in rice seeds, which might represent another strategy to
1014
biofortify Fe/Zn in staple foods (Zhang et al., 2012). However, with respect to Fe, it is
1015
also necessary to increase the biosynthesis of Fe-chelates in the shoot to increase Fe
1016
concentrations in seeds and/or fruits, which rely on Fe supplied via the phloem
1017
(Aoyama et al., 2009, Inoue et al., 2009, Morrissey et al., 2009, Ishimaru et al., 2010,
1018
Kakei et al., 2012).
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1019 1020
Final Remarks 42 Page 42 of 78
The main roles of essential elements have been well documented and they are unlikely
1022
to change. However, many aspects related to whole-plant nutrition needs further
1023
elucidation. For instance, some studies have already described a shift in mineral
1024
nutrition in overexpressing or knockout lines of a certain membrane transporter.
1025
Mutations that increased the content of certain elements in plant tissues usually showed
1026
negative pleiotropic effects in the content of other essential mineral elements and even
1027
on plant growth. It is now widely recognized that a complex network exists between
1028
elements and that biotic and abiotic stress are likely to change not only one element but
1029
the all ionomic profile of a plant. Despite the great advance achieved in last years,
1030
further studies are required to clarify some aspects of plant nutrition. The search for new
1031
players in the field of cation transport still represents a challenge, particularly for trace
1032
essential and beneficial elements. Evidence suggests that the beneficial elements Al, Co
1033
and Na have positive effects on plant growth and stress resistance. Despite that, the
1034
mechanisms behind the beneficial effects of these elements to plants are relatively
1035
underinvestigated, especially when compared to the toxic effects that many of these
1036
elements have at high levels. It is clear that more attention should be given to the effects
1037
of beneficial elements at low levels, since this will not only clarify several basic
1038
processes of plant nutrition but also because fertilizing with beneficial nutrients may
1039
improve crop production and affect plant nutritional value.
1040
This review summarizes the most recent discoveries regarding the function and
1041
regulation of the multiple systems involved in cation transport in plant cells. Since most
1042
of the work has been performed with the model plant Arabidopsis thaliana, the next
1043
step should be the transfer of such knowledge to crop species. The specific properties,
1044
regulation and/or expression patterns of cation transporters/channels in different species
1045
may account for relevant agronomic advances. It is expected that in the upcoming years
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43 Page 43 of 78
many promising results will be achieved in the field of biofortification. However, the
1047
application of biofortification programs will ultimately depend on the cost/benefit
1048
evaluation that farmers, consumers and politics will do. In addition, one of the most
1049
important tasks that researchers need to assure is the careful and systematic analysis of
1050
these foods before they become available to consumers. In this way, it seems that
1051
‘omics’ methods will have a crucial role providing valuable information regarding the
1052
actual safety of these foods.
cr
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1046
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44 Page 44 of 78
1053
Figures Caption
1055
Fig. 1 – Schematic illustration of some of the key transport steps for the uptake of K, Ca
1056
and Mg in the plant (Arabidopsis thaliana was used as model plant). The proteins
1057
identified for mediating some of these transport functions are given in the table below.
1058
The arrows indicate the direction of transport and each letter in the figure corresponds to
1059
the list given in the first column of the table.
1060
Fig. 2 – Fe acquisition strategies in plants: (left) Strategy I in non-graminaceous plants
1061
and (right) Strategy II in graminaceous plants (right). Black boxes represent the
1062
transporters and enzymes that play essential roles in these strategies
1063
Fig. 3 – Schematic illustration of some of the key transport steps for the uptake and
1064
reduction of Cu Mn, Ni and Zn in the plant (Arabidopsis thaliana, Zea mays and
1065
Hordeum vulgare were used as model plants). The proteins identified for mediating
1066
some of these transport/reductive functions are given in the table below. The arrows
1067
indicate the direction of transport/reduction and each letter in the figure corresponds to
1068
the list given in the first column of the table.
1069
Fig. 4 – Tolerance mechanisms for cations in plant cells. Detoxification usually
1070
involves conjugation followed by active sequestration in the vacuole and apoplast,
1071
where the toxic cations can be less harmful. Chelators shown are GSH: glutathione,
1072
MT: metallothioneins, NA: nicotianamine, OA: organic acids, PC: phytochelatins.
1073
Active transporters are shown as black boxes with arrows.
1074
Fig. 5 – Schematic illustration of some of the key transport steps for the translocation of
1075
Fe, Cu Mn and Zn in the plant (Arabidopsis thaliana and Oryza sativa were used as
1076
model plants). The proteins identified for mediating some of these transport functions
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45 Page 45 of 78
are given in the table below. The arrows indicate the direction of transport and each
1078
letter in the figure corresponds to the list given in the first column of the table.
1079
Fig. 6 – Schematic illustration of the subcellular localization of transporters involved in
1080
the movement of K, Ca and Mg in Arabidopsis thaliana. Members of the AtNHX,
1081
AtTPK, AtKCO, AtCHX, AtKEA, AtACA, AtCAX, AtMGT, AtMHX and AtMRS
1082
families are targeted in intracellular membranes: tonoplast (AtNHX1-2, AtACA11,
1083
AtCAX1, AtMGT2, AtMGT3, AtMHX, AtMRS2-11, AtTPK1, AtTPK3, AtTPK5 and
1084
AtKCO3) and chloroplast envelope (AtCHX23 and AtKEA2). The arrows indicate the
1085
direction of transport and each letter in the figure corresponds to the list given in the
1086
first column of the table.
1087
Fig. 7 – Schematic illustration of the subcellular localization of transporters and
1088
reductases involved in Fe, Cu, Mn and Zn movement and reduction, respectively, in
1089
Arabidopsis thaliana and Oryza sativa. Members of the OsVIT, AtFPN, AtNRAMP,
1090
AtBCD, AtPIC, AtFRO, OsMIT, AtCOPT, AtHMA, AtCAX, AtMTP, AtZIP, AtZIF
1091
and AtECA families are targeted in intracellular membranes: vacuole (OsVIT1/2,
1092
AtFPN2, AtCAX2/4-5, AtMTP1/3/11, AtZIF1, AtNRAMP3/4, AtCOPT5 and AtZIP1),
1093
Golgi complex (AtBCD1 and AtECA3), chloroplast (AtPIC1, AtHMA1/6 and
1094
AtFRO7), thylakoid membranes (AtHMA8), mitochondrion (OsMIT and AtFRO3/8)
1095
and endoplasmic reticulum (AtHMA7). The arrows indicate the direction of
1096
transport/reduction and each letter in the figure corresponds to the list given in the first
1097
column of the table.
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1098
46 Page 46 of 78
1098
1101 1102 1103
Ahmad I, Maathuis FJM. Cellular and tissue distribution of potassium: Physiological relevance, mechanisms and regulation. Journal of Plant Physiology. 2014;171:708-14. Ahn SJ, Shin R, Schachtman DP. Expression of KT/KUP genes in arabidopsis and the role
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1100
REFERENCES
of root hairs in K+ uptake. Plant Physiology. 2004;134:1135-45.
cr
1099
Amtmann A, Fischer M, Marsh EL, Stefanovic A, Sanders D, Schachtman DP. The wheat
1105
cDNA LCT1 generates hypersensitivity to sodium in a salt-sensitive yeast strain. Plant
1106
Physiology. 2001;126:1061-71.
us
1104
Ando Y, Nagata S, Yanagisawa S, Yoneyama T. Copper in xylem and phloem saps from
1108
rice (Oryza sativa): the effect of moderate copper concentrations in the growth medium on the
1109
accumulation of five essential metals and a speciation analysis of copper-containing
1110
compounds. Funct Plant Biol. 2013;40:89-100.
M
an
1107
Andres-Colas N, Perea-Garcia A, Puig S, Penaarubia L. Deregulated Copper Transport
1112
Affects Arabidopsis Development Especially in the Absence of Environmental Cycles. Plant
1113
Physiology. 2010;153:170-84.
te
Ac ce p
1114
d
1111
Andres-Colas N, Sancenon V, Rodriguez-Navarro S, Mayo S, Thiele DJ, Ecker JR, et al.
1115
The Arabidopsis heavy metal P-type ATPase HMA5 interacts with metallochaperones and
1116
functions in copper detoxification of roots. Plant Journal. 2006;45:225-36.
1117
Anschutz U, Becker D, Shabala S. Going beyond nutrition: Regulation of potassium
1118
homoeostasis as a common denominator of plant adaptive responses to environment. Journal
1119
of Plant Physiology. 2014;171:670-87.
1120
Aoyama T, Kobayashi T, Takahashi M, Nagasaka S, Usuda K, Kakei Y, et al. OsYSL18 is a
1121
rice iron(III)-deoxymugineic acid transporter specifically expressed in reproductive organs and
1122
phloem of lamina joints. Plant Molecular Biology. 2009;70:681-92.
1123
Apse MP, Blumwald E. Na+ transport in plants. Febs Letters. 2007;581:2247-54. 47 Page 47 of 78
1124
Araki R, Murata J, Murata Y. A Novel Barley Yellow Stripe 1-Like Transporter (HvYSL2)
1125
Localized to the Root Endodermis Transports Metal-Phytosiderophore Complexes. Plant and
1126
Cell Physiology. 2011;52:1931-40. Aranda-Sicilia MN, Cagnac O, Chanroj S, Sze H, Rodriguez-Rosales MP, Venema K.
1128
Arabidopsis KEA2, a homolog of bacterial KefC, encodes a K+/H+ antiporter with a chloroplast
1129
transit peptide. Bba-Biomembranes. 2012;1818:2362-71.
cr
ip t
1127
Ardie SW, Liu S, Takano T. Expression of the AKT1-type K(+) channel gene from
1131
Puccinellia tenuiflora, PutAKT1, enhances salt tolerance in Arabidopsis. Plant Cell Reports.
1132
2010;29:865-74.
us
1130
Arrivault S, Senger T, Kramer U. The Arabidopsis metal tolerance protein AtMTP3
1134
maintains metal homeostasis by mediating Zn exclusion from the shoot under Fe deficiency
1135
and Zn oversupply. Plant Journal. 2006;46:861-79.
M
an
1133
Assunção AGL, Herrero E, Lin YF, Huettel B, Talukdar S, Smaczniak C, et al. Arabidopsis
1137
thaliana transcription factors bZIP19 and bZIP23 regulate the adaptation to zinc deficiency.
1138
Proceedings of the National Academy of Sciences of the United States of America.
1139
2010;107:10296-301.
1141 1142
te
Ac ce p
1140
d
1136
Badri DV, Vivanco JM. Regulation and function of root exudates. Plant Cell and
Environment. 2009;32:666-81.
Barberon M, Zelazny E, Robert S, Conejero G, Curie C, Friml J, et al. Monoubiquitin-
1143
dependent endocytosis of the IRON-REGULATED TRANSPORTER 1 (IRT1) transporter controls
1144
iron uptake in plants. Proceedings of the National Academy of Sciences of the United States of
1145
America. 2011;108:E450-E58.
1146
Barragan V, Leidi EO, Andres Z, Rubio L, De Luca A, Fernandez JA, et al. Ion Exchangers
1147
NHX1 and NHX2 Mediate Active Potassium Uptake into Vacuoles to Regulate Cell Turgor and
1148
Stomatal Function in Arabidopsis. Plant Cell. 2012;24:1127-42.
48 Page 48 of 78
1149 1150
Bashir K, Ishimaru Y, Nishizawa NK. Molecular mechanisms of zinc uptake and translocation in rice. Plant and Soil. 2012;361:189-201. Bashir K, Ishimaru Y, Shimo H, Kakei Y, Senoura T, Takahashi R, et al. Rice phenolics
1152
efflux transporter 2 (PEZ2) plays an important role in solubilizing apoplasmic iron. Soil Sci Plant
1153
Nutr. 2011a;57:803-12.
1155
Bashir K, Ishimaru Y, Shimo H, Nagasaka S, Fujimoto M, Takanashi H, et al. The rice
cr
1154
ip t
1151
mitochondrial iron transporter is essential for plant growth. Nat Commun. 2011b;2.
Bassil E, Ohto MA, Esumi T, Tajima H, Zhu Z, Cagnac O, et al. The Arabidopsis
1157
Intracellular Na+/H+ Antiporters NHX5 and NHX6 Are Endosome Associated and Necessary for
1158
Plant Growth and Development. Plant Cell. 2011;23:224-39.
an
us
1156
Baxter I, Hosmani PS, Rus A, Lahner B, Borevitz JO, Muthukumar B, et al. Root Suberin
1160
Forms an Extracellular Barrier That Affects Water Relations and Mineral Nutrition in
1161
Arabidopsis. Plos Genetics. 2009;5.
d
1163
Benito B, Haro R, Amtmann A, Cuin TA, Dreyer I. The twins K+ and Na+ in plants. Journal of Plant Physiology. 2014;171:723-31.
te
1162
M
1159
Bernal M, Casero D, Singh V, Wilson GT, Grande A, Yang HJ, et al. Transcriptome
1165
Sequencing Identifies SPL7-Regulated Copper Acquisition Genes FRO4/FRO5 and the Copper
1166
Dependence of Iron Homeostasis in Arabidopsis. Plant Cell. 2012;24:738-61.
1167
Ac ce p
1164
Bhatia NP, Walsh KB, Baker AJM. Detection and quantification of ligands involved in
1168
nickel detoxification in a herbaceous Ni hyperaccumulator Stackhousia tryonii Bailey. Journal
1169
of Experimental Botany. 2005;56:1343-49.
1170
Binder BM, Rodriguez FI, Bleecker AB. The Copper Transporter RAN1 Is Essential for
1171
Biogenesis of Ethylene Receptors in Arabidopsis. Journal of Biological Chemistry.
1172
2010;285:37263-70.
49 Page 49 of 78
1173
Cailliatte R, Schikora A, Briat JF, Mari S, Curie C. High-Affinity Manganese Uptake by the
1174
Metal Transporter NRAMP1 Is Essential for Arabidopsis Growth in Low Manganese Conditions.
1175
Plant Cell. 2010;22:904-17. Catty P, Boutigny S, Miras R, Joyard J, Rolland N, Seigneurin-Berny D. Biochemical
1177
Characterization of AtHMA6/PAA1, a Chloroplast Envelope Cu(I)-ATPase. Journal of Biological
1178
Chemistry. 2011;286:36188-97.
cr
ip t
1176
Cellier F, Conejero G, Ricaud L, Luu DT, Lepetit M, Gosti F, et al. Characterization of
1180
AtCHX17, a member of the cation/H+ exchangers, CHX family, from Arabidopsis thaliana
1181
suggests a role in K+ homeostasis. Plant Journal. 2004;39:834-46.
us
1179
Centofanti T, Sayers Z, Cabello-Conejo MI, Kidd P, Nishizawa NK, Kakei Y, et al. Xylem
1183
exudate composition and root-to-shoot nickel translocation in Alyssum species. Plant and Soil.
1184
2013;373:59-75.
1186
M
Chen J, Li LG, Liu ZH, Yuan YJ, Guo LL, Mao DD, et al. Magnesium transporter AtMGT9 is essential for pollen development in Arabidopsis. Cell Research. 2009;19:887-98.
d
1185
an
1182
Cheng NH, Pittman JK, Shigaki T, Lachmansingh J, LeClere S, Lahner B, et al. Functional
1188
association of Arabidopsis CAX1 and CAX3 is required for normal growth and ion homeostasis.
1189
Plant Physiology. 2005;138:2048-60.
Ac ce p
1190
te
1187
Chu HH, Chiecko J, Punshon T, Lanzirotti A, Lahner B, Salt DE, et al. Successful
1191
Reproduction Requires the Function of Arabidopsis YELLOW STRIPE-LIKE1 and YELLOW STRIPE-
1192
LIKE3 Metal-Nicotianamine Transporters in Both Vegetative and Reproductive Structures. Plant
1193
Physiology. 2010;154:197-210.
1194
Conn SJ, Conn V, Tyerman SD, Kaiser BN, Leigh RA, Gilliham M. Magnesium
1195
transporters, MGT2/MRS2-1 and MGT3/MRS2-5, are important for magnesium partitioning
1196
within Arabidopsis thaliana mesophyll vacuoles. New Phytologist. 2011a;190:583-94.
50 Page 50 of 78
1197
Conn SJ, Gilliham M, Athman A, Schreiber AW, Baumann U, Moller I, et al. Cell-Specific
1198
Vacuolar Calcium Storage Mediated by CAX1 Regulates Apoplastic Calcium Concentration, Gas
1199
Exchange, and Plant Productivity in Arabidopsis. Plant Cell. 2011b;23:240-57. Connorton JM, Webster RE, Cheng NH, Pittman JK. Knockout of Multiple Arabidopsis
1201
Cation/H+ Exchangers Suggests Isoform-Specific Roles in Metal Stress Response, Germination
1202
and Seed Mineral Nutrition. Plos One. 2012;7.
cr
1204
Cosio C, DeSantis L, Frey B, Diallo S, Keller C. Distribution of cadmium in leaves of Thiaspi caerulescens. Journal of Experimental Botany. 2005;56:765-75.
us
1203
ip t
1200
Davenport RJ, Munoz-Mayor A, Jha D, Essah PA, Rus A, Tester M. The Na+ transporter
1206
AtHKT1;1 controls retrieval of Na+ from the xylem in Arabidopsis. Plant Cell and Environment.
1207
2007;30:497-507.
1209
Dayod M, Tyerman SD, Leigh RA, Gilliham M. Calcium storage in plants and the
M
1208
an
1205
implications for calcium biofortification. Protoplasma. 2010;247:215-31. Deinlein U, Weber M, Schmidt H, Rensch S, Trampczynska A, Hansen TH, et al. Elevated
1211
Nicotianamine Levels in Arabidopsis halleri Roots Play a Key Role in Zinc Hyperaccumulation.
1212
Plant Cell. 2012;24:708-23.
te
Ac ce p
1213
d
1210
del Pozo T, Cambiazo V, Gonzalez M. Gene expression profiling analysis of copper
1214
homeostasis in Arabidopsis thaliana. Biochemical and Biophysical Research Communications.
1215
2010;393:248-52.
1216
Delhaize E, Gruber BD, Pittman JK, White RG, Leung H, Miao YS, et al. A role for the
1217
AtMTP11 gene of Arabidopsis in manganese transport and tolerance. Plant Journal.
1218
2007;51:198-210.
1219 1220
Demidchik V, Maathuis FJM. Physiological roles of nonselective cation channels in plants: from salt stress to signalling and development. New Phytologist. 2007;175:387-404.
51 Page 51 of 78
1221
Deng W, Luo KM, Li DM, Zheng XL, Wei XY, Smith W, et al. Overexpression of an
1222
Arabidopsis magnesium transport gene, AtMGT1, in Nicotiana benthamiana confers Al
1223
tolerance. Journal of Experimental Botany. 2006;57:4235-43. Drummond RSM, Tutone A, Li YC, Gardner RC. A putative magnesium transporter
1225
AtMRS2-11 is localized to the plant chloroplast envelope membrane system. Plant Science.
1226
2006;170:78-89.
cr
1227
Durrett TP, Gassmann W, Rogers EE. The FRD3-mediated efflux of citrate into the root vasculature is necessary for efficient iron translocation. Plant Physiology. 2007;144:197-205.
us
1228
ip t
1224
Duy D, Stube R, Wanner G, Philippar K. The Chloroplast Permease PIC1 Regulates Plant
1230
Growth and Development by Directing Homeostasis and Transport of Iron. Plant Physiology.
1231
2011;155:1709-22.
an
1229
Edmond C, Shigaki T, Ewert S, Nelson MD, Connorton JM, Chalova V, et al. Comparative
1233
analysis of CAX2-like cation transporters indicates functional and regulatory diversity. Biochem
1234
J. 2009;418:145-54.
d
M
1232
Elbaz B, Shoshani-Knaani N, David-Assael O, Mizrachy-Dagri T, Mizrahi K, Saul H, et al.
1236
High expression in leaves of the zinc hyperaccumulator Arabidopsis halleri of AhMHX, a
1237
homolog of an Arabidopsis thaliana vacuolar metal/proton exchanger. Plant Cell and
1238
Environment. 2006;29:1179-90.
Ac ce p
1239
te
1235
Fedotov PS, Kordel W, Miro M, Peijnenburg WJGM, Wennrich R, Huang PM. Extraction
1240
and Fractionation Methods for Exposure Assessment of Trace Metals, Metalloids, and
1241
Hazardous Organic Compounds in Terrestrial Environments. Crit Rev Env Sci Tec.
1242
2012;42:1117-71.
1243 1244
FNB/IOM. PART III: VITAMINS AND MINERALS.
Essential Guide to Nutrient Requirements: The National Academies Press; 2006. p. 167-422.
1245 1246
Dietary Reference Intakes: The
Forde BG. Glutamate signalling in roots. Journal of Experimental Botany. 2014;65:77987. 52 Page 52 of 78
1247 1248
Fox TC, Guerinot ML. MOLECULAR BIOLOGY OF CATION TRANSPORT IN PLANTS. Annu Rev Plant Phys. 1998;49:669-96. Fukao Y, Ferjani A, Tomioka R, Nagasaki N, Kurata R, Nishimori Y, et al. iTRAQ Analysis
1250
Reveals Mechanisms of Growth Defects Due to Excess Zinc in Arabidopsis. Plant Physiology.
1251
2011;155:1893-907.
ip t
1249
Fulgenzi FR, Peralta ML, Mangano S, Danna CH, Vallejo AJ, Puigdomenech P, et al. The
1253
ionic environment controls the contribution of the barley HvHAK1 transporter to potassium
1254
acquisition. Plant Physiology. 2008;147:252-62.
us
cr
1252
Gajdanowicz P, Michard E, Sandmann M, Rocha M, Correa LGG, Ramirez-Aguilar SJ, et
1256
al. Potassium (K+ ) gradients serve as a mobile energy source in plant vascular tissues.
1257
Proceedings of the National Academy of Sciences of the United States of America.
1258
2011;108:864-69.
M
an
1255
Garcia-Molina A, Andres-Colas N, Perea-Garcia A, del Valle-Tascon S, Penarrubia L, Puig
1260
S. The intracellular Arabidopsis COPT5 transport protein is required for photosynthetic
1261
electron transport under severe copper deficiency. Plant Journal. 2011;65:848-60.
te
d
1259
Gebert M, Meschenmoser K, Svidova S, Weghuber J, Schweyen R, Eifler K, et al. A Root-
1263
Expressed Magnesium Transporter of the MRS2/MGT Gene Family in Arabidopsis thaliana
1264
Allows for Growth in Low-Mg2+ Environments. Plant Cell. 2009;21:4018-30.
1265
Ac ce p
1262
Geiger D, Becker D, Vosloh D, Gambale F, Palme K, Rehers M, et al. Heteromeric
1266
AtKC1.AKT1 Channels in Arabidopsis Roots Facilitate Growth under K+-limiting Conditions.
1267
Journal of Biological Chemistry. 2009;284:21288-95.
1268
Gendre D, Czernic P, Conejero G, Pianelli K, Briat JF, Lebrun M, et al. TcYSL3, a member
1269
of the YSL gene family from the hyper-accumulator Thlaspi caerulescens, encodes a
1270
nicotianamine-Ni/Fe transporter. Plant Journal. 2007;49:1-15.
53 Page 53 of 78
1271
Ghasemi R, Ghaderian SM, Kramer U. Interference of nickel with copper and iron
1272
homeostasis contributes to metal toxicity symptoms in the nickel hyperaccumulator plant
1273
Alyssum inflatum. New Phytologist. 2009;184:566-80. Guo WJ, Meetam M, Goldsbrough PB. Examining the specific contributions of
1275
individual Arabidopsis metallothioneins to copper distribution and metal tolerance. Plant
1276
Physiology. 2008;146:1697-706.
cr
ip t
1274
Gustin JL, Loureiro ME, Kim D, Na G, Tikhonova M, Salt DE. MTP1-dependent Zn
1278
sequestration into shoot vacuoles suggests dual roles in Zn tolerance and accumulation in Zn-
1279
hyperaccumulating plants. Plant Journal. 2009;57:1116-27.
1284 1285 1286 1287 1288
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M
1283
Haferkamp I, Linka N. Functional expression and characterisation of membrane transport proteins. Plant Biology. 2012;14:675-90.
Hamamoto S, Uozumi N. Organelle-localized potassium transport systems in plants.
d
1282
facilitator family of ion transporters. Bmc Evol Biol. 2011;11.
Journal of Plant Physiology. 2014;171:743-47.
te
1281
Gustin JL, Zanis MJ, Salt DE. Structure and evolution of the plant cation diffusion
Hänsch R, Mendel RR. Physiological functions of mineral micronutrients (Cu, Zn, Mn,
Ac ce p
1280
us
1277
Fe, Ni, Mo, B, Cl). Curr Opin Plant Biol. 2009;12:259-66. Haro R, Banuelos MA, Senn MAE, Barrero-Gil J, Rodriguez-Navarro A. HKT1 mediates
1289
sodium uniport in roots. Pitfalls in the expression of HKT1 in yeast. Plant Physiology.
1290
2005;139:1495-506.
1291 1292
Harris WR, Sammons RD, Grabiak RC. A speciation model of essential trace metal ions
in phloem. J Inorg Biochem. 2012;116:140-50.
1293
Haydon MJ, Kawachi M, Wirtz M, Hillmer S, Hell R, Kramer U. Vacuolar Nicotianamine
1294
Has Critical and Distinct Roles under Iron Deficiency and for Zinc Sequestration in Arabidopsis.
1295
Plant Cell. 2012;24:724-37.
1296
Hedrich R, Marten I. TPC1-SV Channels Gain Shape. Molecular Plant. 2011;4:428-41. 54 Page 54 of 78
1297 1298
Hinsinger P, Bengough AG, Vetterlein D, Young IM. Rhizosphere: biophysics, biogeochemistry and ecological relevance. Plant and Soil. 2009;321:117-52. Hirschi KD. Nutrient Biofortification of Food Crops. Annu Rev Nutr. 2009;29:401-21.
1300
Horie T, Sugawara M, Okada T, Taira K, Kaothien-Nakayama P, Katsuhara M, et al. Rice
1301
sodium-insensitive potassium transporter, OsHAK5, confers increased salt tolerance in tobacco
1302
BY2 cells. Journal of bioscience and bioengineering. 2011;111:346-56.
cr
ip t
1299
Hu YT, Ming F, Chen WW, Yan JY, Xu ZY, Li GX, et al. TcOPT3, a Member of Oligopeptide
1304
Transporters from the Hyperaccumulator Thlaspi caerulescens, Is a Novel Fe/Zn/Cd/Cu
1305
Transporter. Plos One. 2012;7.
us
1303
Inoue H, Kobayashi T, Nozoye T, Takahashi M, Kakei Y, Suzuki K, et al. Rice OsYSL15 Is
1307
an Iron-regulated Iron(III)-Deoxymugineic Acid Transporter Expressed in the Roots and Is
1308
Essential for Iron Uptake in Early Growth of the Seedlings. Journal of Biological Chemistry.
1309
2009;284:3470-79.
M
an
1306
Irtelli B, Petrucci WA, Navari-Izzo F. Nicotianamine and histidine/proline are,
1311
respectively, the most important copper chelators in xylem sap of Brassica carinata under
1312
conditions of copper deficiency and excess. Journal of Experimental Botany. 2009;60:269-77.
te
Ac ce p
1313
d
1310
Ishimaru Y, Kakei Y, Shimo H, Bashir K, Sato Y, Sato Y, et al. A Rice Phenolic Efflux
1314
Transporter Is Essential for Solubilizing Precipitated Apoplasmic Iron in the Plant Stele. Journal
1315
of Biological Chemistry. 2011;286:24649-55.
1316
Ishimaru Y, Masuda H, Bashir K, Inoue H, Tsukamoto T, Takahashi M, et al. Rice metal-
1317
nicotianamine transporter, OsYSL2, is required for the long-distance transport of iron and
1318
manganese. Plant Journal. 2010;62:379-90.
1319
Ishimaru Y, Takahashi R, Bashir K, Shimo H, Senoura T, Sugimoto K, et al. Characterizing
1320
the role of rice NRAMP5 in Manganese, Iron and Cadmium Transport. Scientific Reports.
1321
2012;2.
55 Page 55 of 78
1322
Jabnoune M, Espeout S, Mieulet D, Fizames C, Verdeil JL, Conejero G, et al. Diversity in
1323
Expression Patterns and Functional Properties in the Rice HKT Transporter Family. Plant
1324
Physiology. 2009;150:1955-71.
1327 1328
ip t
1326
Jeanguenin L, Alcon C, Duby G, Boeglin M, Cherel I, Gaillard I, et al. AtKC1 is a general modulator of Arabidopsis inward Shaker channel activity. The Plant Journal. 2011;67:570-82.
Jeong J, Connolly EL. Iron uptake mechanisms in plants: Functions of the FRO family of
cr
1325
ferric reductases. Plant Science. 2009;176:709-14.
Jeong J, Guerinot ML. Biofortified and bioavailable: The gold standard, for plant-based
1330
diets. Proceedings of the National Academy of Sciences of the United States of America.
1331
2008;105:1777-78.
an
us
1329
Johansson I, Wulfetange K, Poree F, Michard E, Gajdanowicz P, Lacombe B, et al.
1333
External K+ modulates the activity of the Arabidopsis potassium channel SKOR via an unusual
1334
mechanism. Plant Journal. 2006;46:269-81.
d
1336
Jones B, Ljung K. Subterranean space exploration: the development of root system architecture. Curr Opin Plant Biol. 2012;15:97-102.
te
1335
M
1332
Jung HI, Gayomba SR, Rutzke MA, Craft E, Kochian LV, Vatamaniuk OK. COPT6 Is a
1338
Plasma Membrane Transporter That Functions in Copper Homeostasis in Arabidopsis and Is a
1339
Novel Target of SQUAMOSA Promoter-binding Protein-like 7. Journal of Biological Chemistry.
1340
2012;287:33252-67.
1341 1342 1343 1344 1345 1346
Ac ce p
1337
Kabata-Pendias A. Soil–plant transfer of trace elements—an environmental issue.
Geoderma. 2004;122:143-49.
Kakei Y, Ishimaru Y, Kobayashi T, Yamakawa T, Nakanishi H, Nishizawa NK. OsYSL16 plays a role in the allocation of iron. Plant Molecular Biology. 2012;79:583-94. Karley AJ, White PJ. Moving cationic minerals to edible tissues: potassium, magnesium, calcium. Curr Opin Plant Biol. 2009;12:291-98.
56 Page 56 of 78
1347
Kawachi M, Kobae Y, Mori H, Tomioka R, Lee Y, Maeshima M. A Mutant Strain
1348
Arabidopsis thaliana that Lacks Vacuolar Membrane Zinc Transporter MTP1 Revealed the
1349
Latent Tolerance to Excessive Zinc. Plant and Cell Physiology. 2009;50:1156-70. Kim CK, Han JS, Lee HS, Oh JY, Shigaki T, Park SH, et al. Expression of an Arabidopsis
1351
CAX2 variant in potato tubers increases calcium levels with no accumulation of manganese.
1352
Plant Cell Reports. 2006a;25:1226-32.
cr
ip t
1350
Kim SA, Punshon T, Lanzirotti A, Li LT, Alonso JM, Ecker JR, et al. Localization of iron in
1354
Arabidopsis seed requires the vacuolar membrane transporter VIT1. Science. 2006b;314:1295-
1355
98.
1357
Kim YY, Choi H, Segami S, Cho HT, Martinoia E, Maeshima M, et al. AtHMA1
an
1356
us
1353
contributes to the detoxification of excess Zn(II) in Arabidopsis. Plant Journal. 2009;58:737-53. Klaumann S, Nickolaus SD, Furst SH, Starck S, Schneider S, Neuhaus HE, et al. The
1359
tonoplast copper transporter COPT5 acts as an exporter and is required for interorgan
1360
allocation of copper in Arabidopsis thaliana. New Phytologist. 2011;192:393-404.
1363
d
te
1362
Kobayashi T, Nishizawa NK. Iron Uptake, Translocation, and Regulation in Higher Plants. Annual Review of Plant Biology. 2012;63:131-52.
Ac ce p
1361
M
1358
Krebs M, Beyhl D, Gorlich E, Al-Rasheid KAS, Marten I, Stierhof YD, et al. Arabidopsis V-
1364
ATPase activity at the tonoplast is required for efficient nutrient storage but not for sodium
1365
accumulation. Proceedings of the National Academy of Sciences of the United States of
1366
America. 2010;107:3251-56.
1367 1368
Kronzucker HJ, Britto DT. Sodium transport in plants: a critical review. New Phytologist.
2011;189:54-81.
1369
Lanquar V, Lelievre F, Bolte S, Hames C, Alcon C, Neumann D, et al. Mobilization of
1370
vacuolar iron by AtNRAMP3 and AtNRAMP4 is essential for seed germination on low iron.
1371
Embo J. 2005;24:4041-51.
57 Page 57 of 78
Lanquar V, Ramos MS, Lelievre F, Barbier-Brygoo H, Krieger-Liszkay A, Kramer U, et al.
1373
Export of Vacuolar Manganese by AtNRAMP3 and AtNRAMP4 Is Required for Optimal
1374
Photosynthesis and Growth under Manganese Deficiency. Plant Physiology. 2010;152:1986-99.
1375
Larbi A, Morales F, Abadia A, Abadia J. Changes in iron and organic acid concentrations
1376
in xylem sap and apoplastic fluid of iron-deficient Beta vulgaris plants in response to iron
1377
resupply. Journal of Plant Physiology. 2010;167:255-60.
cr
ip t
1372
Larras F, Regier N, Planchon S, Pote J, Renaut J, Cosio C. Physiological and proteomic
1379
changes suggest an important role of cell walls in the high tolerance to metals of Elodea
1380
nuttallii. Journal of Hazardous Materials. 2013;263:575-83.
1383 1384
an
1382
Lee S, An G. Over-expression of OsIRT1 leads to increased iron and zinc accumulations in rice. Plant Cell and Environment. 2009;32:408-16.
Lee S, Kim SA, Lee J, Guerinot ML, An G. Zinc Deficiency-Inducible OsZIP8 Encodes a
M
1381
us
1378
Plasma Membrane-Localized Zinc Transporter in Rice. Molecules and Cells. 2010;29:551-58. Lee SM, Kim HS, Han HJ, Moon BC, Kim CY, Harper JF, et al. Identification of a
1386
calmodulin-regulated autoinhibited Ca2+-ATPase (ACA11) that is localized to vacuole
1387
membranes in Arabidopsis. Febs Letters. 2007;581:3943-49.
te
Ac ce p
1388
d
1385
Li XY, Chanroj S, Wu ZY, Romanowsky SM, Harper JF, Sze H. A distinct endosomal
1389
Ca2+/Mn2+ pump affects root growth through the secretory process. Plant Physiology.
1390
2008;147:1675-89.
1391
Lin YF, Liang HM, Yang SY, Boch A, Clemens S, Chen CC, et al. Arabidopsis IRT3 is a zinc-
1392
regulated and plasma membrane localized zinc/iron transporter. New Phytologist.
1393
2009;182:392-404.
1394 1395 1396 1397
Ma W, Smigel A, Verma R, Berkowitz GA. Cyclic nucleotide gated channels and related signaling components in plant innate immunity. Plant Signaling & Behavior. 2009;4:277-82. Maathuis FJM. Physiological functions of mineral macronutrients. Curr Opin Plant Biol. 2009;12:250-58. 58 Page 58 of 78
1398 1399
Manohar M, Shigaki T, Hirschi KD. Plant cation/H+ exchangers (CAXs): biological functions and genetic manipulations. Plant Biology. 2011;13:561-69. Mao DD, Tian LF, Li LG, Chen J, Deng PY, Li DP, et al. AtMGT7: An Arabidopsis Gene
1401
Encoding a Low-Affinity Magnesium Transporter. Journal of Integrative Plant Biology.
1402
2008;50:1530-38.
ip t
1400
Mari S, Gendre D, Pianelli K, Ouerdane L, Lobinski R, Briat JF, et al. Root-to-shoot long-
1404
distance circulation of nicotianamine and nicotianamine-nickel chelates in the metal
1405
hyperaccumulator Thlaspi caerulescens. Journal of Experimental Botany. 2006;57:4111-22.
1408 1409
us
Context. Annual Review of Plant Biology. 2012;63:183-213.
an
1407
Martinoia E, Meyer S, De Angeli A, Nagy R. Vacuolar Transporters in Their Physiological
McAinsh MR, Pittman JK. Shaping the calcium signature. New Phytologist. 2009;181:275-94.
M
1406
cr
1403
Mian A, Oomen RJFJ, Isayenkov S, Sentenac H, Maathuis FJM, Very AA. Over-
1411
expression of an Na+- and K+-permeable HKT transporter in barley improves salt tolerance.
1412
Plant Journal. 2011;68:468-79.
te
d
1410
Mikkelsen MD, Pedas P, Schiller M, Vincze E, Mills RF, Borg S, et al. Barley HvHMA1 Is a
1414
Heavy Metal Pump Involved in Mobilizing Organellar Zn and Cu and Plays a Role in Metal
1415
Loading into Grains. Plos One. 2012;7.
1416
Ac ce p
1413
Millaleo R, Reyes-Diaz M, Alberdi M, Ivanov AG, Krol M, Huner NPA. Excess manganese
1417
differentially inhibits photosystem I versus II in Arabidopsis thaliana. Journal of Experimental
1418
Botany. 2013;64:343-54.
1419
Mills RF, Doherty ML, Lopez-Marques RL, Weimar T, Dupree P, Palmgren MG, et al.
1420
ECA3, a Golgi-localized P-2A-type ATPase, plays a crucial role in manganese nutrition in
1421
Arabidopsis. Plant Physiology. 2008;146:116-28.
1422 1423
Mills RF, Peaston KA, Runions J, Williams LE. HvHMA2, a P-1B-ATPase from Barley, Is Highly Conserved among Cereals and Functions in Zn and Cd Transport. Plos One. 2012;7. 59 Page 59 of 78
1424
Milner MJ, Seamon J, Craft E, Kochian LV. Transport properties of members of the ZIP
1425
family in plants and their role in Zn and Mn homeostasis. Journal of Experimental Botany.
1426
2013;64:369-81. Moller IS, Gilliham M, Jha D, Mayo GM, Roy SJ, Coates JC, et al. Shoot Na+ Exclusion
1428
and Increased Salinity Tolerance Engineered by Cell Type-Specific Alteration of Na+ Transport
1429
in Arabidopsis. Plant Cell. 2009;21:2163-78.
cr
ip t
1427
Morel M, Crouzet J, Gravot A, Auroy P, Leonhardt N, Vavasseur A, et al. AtHMA3, a P-
1431
1B-ATPase Allowing Cd/Zn/Co/Pb Vacuolar Storage in Arabidopsis. Plant Physiology.
1432
2009;149:894-904.
us
1430
Moreno-Risueno MA, Van Norman JM, Moreno A, Zhang JY, Ahnert SE, Benfey PN.
1434
Oscillating Gene Expression Determines Competence for Periodic Arabidopsis Root Branching.
1435
Science. 2010;329:1306-11.
1437
M
Morrissey J, Baxter IR, Lee J, Li LT, Lahner B, Grotz N, et al. The Ferroportin Metal Efflux Proteins Function in Iron and Cobalt Homeostasis in Arabidopsis. Plant Cell. 2009;21:3326-38.
d
1436
an
1433
Mortimer JC, Laohavisit A, Macpherson N, Webb A, Brownlee C, Battey NH, et al.
1439
Annexins: multifunctional components of growth and adaptation. Journal of Experimental
1440
Botany. 2008;59:533-44.
Ac ce p
1441
te
1438
Mukherjee I, Campbell NH, Ash JS, Connolly EL. Expression profiling of the Arabidopsis
1442
ferric chelate reductase (FRO) gene family reveals differential regulation by iron and copper.
1443
Planta. 2006;223:1178-90.
1444
Murata Y, Harada E, Sugase K, Namba K, Horikawa M, Ma JF, et al. Specific transporter
1445
for iron(III)-phytosiderophore complex involved in iron uptake by barley roots. Pure and
1446
Applied Chemistry. 2008;80:2689-97.
1447
Nieves-Cordones M, Aleman F, Martinez V, Rubio F. The Arabidopsis thaliana HAK5 K+
1448
Transporter Is Required for Plant Growth and K+ Acquisition from Low K+ Solutions under
1449
Saline Conditions. Molecular Plant. 2010;3:326-33. 60 Page 60 of 78
1450
Nieves-Cordones M, Aleman F, Martinez V, Rubio F. K+ uptake in plant roots. The
1451
systems involved, their regulation and parallels in other organisms. Journal of Plant Physiology.
1452
2014;171:688-95.
1454
Nishida S, Aisu A, Mizuno T. Induction of IRT1 by the nickel-induced iron-deficient response in Arabidopsis. Plant Signaling & Behavior. 2012;7:329-31.
ip t
1453
Nishiyama R, Kato M, Nagata S, Yanagisawa S, Yoneyama T. Identification of Zn-
1456
Nicotianamine and Fe-2'-Deoxymugineic Acid in the Phloem Sap from Rice Plants (Oryza sativa
1457
L.). Plant and Cell Physiology. 2012;53:381-90.
us
cr
1455
Nozoye T, Nagasaka S, Kobayashi T, Takahashi M, Sato Y, Sato Y, et al.
1459
Phytosiderophore Efflux Transporters Are Crucial for Iron Acquisition in Graminaceous Plants.
1460
Journal of Biological Chemistry. 2011;286:5446-54.
1462
Nozoye T, Nakanishi H, Nishizawa NK. Characterizing the Crucial Components of Iron
M
1461
an
1458
Homeostasis in the Maize Mutants ys1 and ys3. Plos One. 2013;8. Oomen RJFJ, Wu J, Lelievre F, Blanchet S, Richaud P, Barbier-Brygoo H, et al. Functional
1464
characterization of NRAMP3 and NRAMP4 from the metal hyperaccumulator Thlaspi
1465
caerulescens. New Phytologist. 2009;181:637-50.
1467 1468 1469 1470 1471 1472 1473 1474 1475
te
Ac ce p
1466
d
1463
Pal R, Rai JPN. Phytochelatins: Peptides Involved in Heavy Metal Detoxification. Appl
Biochem Biotech. 2010;160:945-63. Palm C, Sanchez P, Ahamed S, Awiti A. Soils: A contemporary perspective. Annu Rev
Env Resour. 2007;32:99-129.
Palmgren MG. Plant plasma membrane H+-ATPases: Powerhouses for nutrient uptake.
Annu Rev Plant Phys. 2001a;52:817-45. Palmgren MG. PLANT PLASMA MEMBRANE H+-ATPases: Powerhouses for Nutrient Uptake. Annu Rev Plant Phys. 2001b;52:817-45. Park S, Kang TS, Kim CK, Han JS, Kim S, Smith RH, et al. Genetic manipulation for enhancing calcium content in potato tuber. J Agr Food Chem. 2005;53:5598-603. 61 Page 61 of 78
1476 1477
Park S, Kim CK, Pike LM, Smith RH, Hirschi KD. Increased calcium in carrots by expression of an Arabidopsis H+/Ca2+ transporter. Mol Breeding. 2004;14:275-82. Pedas P, Ytting CK, Fuglsang AT, Jahn TP, Schjoerring JK, Husted S. Manganese
1479
efficiency in barley: Identification and characterization of the metal ion transporter HvIRT1.
1480
Plant Physiology. 2008;148:455-66.
ip t
1478
Petersson SV, Johansson AI, Kowalczyk M, Makoveychuk A, Wang JY, Moritz T, et al. An
1482
Auxin Gradient and Maximum in the Arabidopsis Root Apex Shown by High-Resolution Cell-
1483
Specific Analysis of IAA Distribution and Synthesis. Plant Cell. 2009;21:1659-68.
1486 1487
us
of beneficial elements. Curr Opin Plant Biol. 2009;12:267-74.
an
1485
Pilon-Smits EAH, Quinn CF, Tapken W, Malagoli M, Schiavon M. Physiological functions
Pilon M, Cohu CM, Ravet K, Abdel-Ghany SE, Gaymard F. Essential transition metal homeostasis in plants. Curr Opin Plant Biol. 2009;12:347-57.
M
1484
cr
1481
Pineau C, Loubet S, Lefoulon C, Chalies C, Fizames C, Lacombe B, et al. Natural
1489
Variation at the FRD3 MATE Transporter Locus Reveals Cross-Talk between Fe Homeostasis
1490
and Zn Tolerance in Arabidopsis thaliana. Plos Genetics. 2012;8.
1492 1493
te
Pinto E, Aguiar AARM, Ferreira IMPLVO. Influence of Soil Chemistry and Plant
Ac ce p
1491
d
1488
Physiology in the Phytoremediation of Cu, Mn, and Zn. Crit Rev Plant Sci. 2014a;33:351-73. Pinto E, Almeida AA, Aguiar AARM, Ferreira IMPLVO. Changes in macrominerals, trace
1494
elements and pigments content during lettuce (Lactuca sativa L.) growth: Influence of soil
1495
composition. Food Chem. 2014b;152:603-11.
1496
Pinto E, Almeida AA, Ferreira IMPLVO. Assessment of metal(loid)s phytoavailability in
1497
intensive agricultural soils by the application of single extractions to rhizosphere soil.
1498
Ecotoxicology and Environmental Safety. 2015;113:418-24.
1499 1500 1501
Pittman JK. Managing the manganese: molecular mechanisms of manganese transport and homeostasis. New Phytologist. 2005;167:733-42. Pittman JK. Vacuolar Ca2+ uptake. Cell Calcium. 2011;50:139-46. 62 Page 62 of 78
1502
Pottosin I, Dobrovinskaya O. Non-selective cation channels in plasma and vacuolar
1503
membranes and their contribution to K+ transport. Journal of Plant Physiology. 2014;171:732-
1504
42.
1506
Puig S, Penarrubia L. Placing metal micronutrients in context: transport and
ip t
1505
distribution in plants. Curr Opin Plant Biol. 2009;12:299-306.
Ranf S, Wunnenberg P, Lee J, Becker D, Dunkel M, Hedrich R, et al. Loss of the vacuolar
1508
cation channel, AtTPC1, does not impair Ca2+ signals induced by abiotic and biotic stresses.
1509
Plant Journal. 2008;53:287-99.
us
cr
1507
Rellan-Alvarez R, Abadia J, Alvarez-Fernandez A. Formation of metal-nicotianamine
1511
complexes as affected by pH, ligand exchange with citrate and metal exchange. A study by
1512
electrospray ionization time-of-flight mass spectrometry. Rapid Commun Mass Sp.
1513
2008;22:1553-62.
M
an
1510
Rellan-Alvarez R, Giner-Martinez-Sierra J, Orduna J, Orera I, Rodriguez-Castrillon JA,
1515
Garcia-Alonso JI, et al. Identification of a Tri-Iron(III), Tri-Citrate Complex in the Xylem Sap of
1516
Iron-Deficient Tomato Resupplied with Iron: New Insights into Plant Iron Long-Distance
1517
Transport. Plant and Cell Physiology. 2010;51:91-102.
te
Ac ce p
1518
d
1514
Richau KH, Kozhevnikova AD, Seregin IV, Vooijs R, Koevoets PLM, Smith JAC, et al.
1519
Chelation by histidine inhibits the vacuolar sequestration of nickel in roots of the
1520
hyperaccumulator Thlaspi caerulescens. New Phytologist. 2009;183:106-16.
1521
Rubio F, Nieves-Cordones M, Aleman F, Martinez V. Relative contribution of AtHAK5
1522
and AtAKT1 to K(+) uptake in the high-affinity range of concentrations. Physiologia Plantarum.
1523
2008;134:598-608.
1524
Saito A, Saito M, Ichikawa Y, Yoshiba M, Tadano T, Miwa E, et al. Difference in the
1525
distribution and speciation of cellular nickel between nickel-tolerant and non-tolerant
1526
Nicotiana tabacum L. cv. BY-2 cells. Plant Cell and Environment. 2010;33:174-87.
63 Page 63 of 78
1527
Sancenon V, Puig S, Mateu-Andres I, Dorcey E, Thiele DJ, Penarrubia L. The Arabidopsis
1528
copper transporter COPT1 functions in root elongation and pollen development. Journal of
1529
Biological Chemistry. 2004;279:15348-55. Sanchez-Pardo B, Fernandez-Pascual M, Zornoza P. Copper microlocalisation,
1531
ultrastructural alterations and antioxidant responses in the nodules of white lupin and soybean
1532
plants grown under conditions of copper excess. Environmental and Experimental Botany.
1533
2012;84:52-60.
1537 1538 1539
cr
us
1536
roots. New Phytologist. 2009;183:1072-84.
Sasaki A, Yamaji N, Xia JX, Ma JF. OsYSL6 Is Involved in the Detoxification of Excess
an
1535
Santi S, Schmidt W. Dissecting iron deficiency-induced proton extrusion in Arabidopsis
Manganese in Rice. Plant Physiology. 2011;157:1832-40.
Sassi A, Mieulet D, Khan I, Moreau B, Gaillard I, Sentenac H, et al. The Rice Monovalent
M
1534
ip t
1530
Cation Transporter OsHKT2;4: Revisited Ionic Selectivity. Plant Physiology. 2012;160:498-510. Schaaf G, Honsbein A, Meda AR, Kirchner S, Wipf D, von Wiren N. AtIREG2 encodes a
1541
tonoplast transport protein involved in iron-dependent nickel detoxification in Arabidopsis
1542
thaliana roots. Journal of Biological Chemistry. 2006;281:25532-40.
te
Ac ce p
1543
d
1540
Seo PJ, Park J, Park MJ, Kim YS, Kim SG, Jung JH, et al. A Golgi-localized MATE
1544
transporter mediates iron homoeostasis under osmotic stress in Arabidopsis. Biochem J.
1545
2012;442:551-61.
1546
Shanmugam V, Lo JC, Wu CL, Wang SL, Lai CC, Connolly EL, et al. Differential expression
1547
and regulation of iron-regulated metal transporters in Arabidopsis halleri and Arabidopsis
1548
thaliana - the role in zinc tolerance. New Phytologist. 2011;190:125-37.
1549 1550 1551 1552
Shaul O. Magnesium transport and function in plants: the tip of the iceberg. Biometals. 2002;15:309-23. Shaul O, Hilgemann DW, de-Almeida-Engler J, Van Montagu M, Inzé D, Galili G. Cloning and characterization of a novel Mg2+/H+ exchanger. The EMBO Journal. 1999;18:3973-80. 64 Page 64 of 78
1553 1554
Shi HZ, Quintero FJ, Pardo JM, Zhu JK. The putative plasma membrane Na+/H+ antiporter SOS1 controls long-distance Na+ transport in plants. Plant Cell. 2002;14:465-77. Siemianowski O, Barabasz A, Kendziorek M, Ruszczynska A, Bulska E, Williams LE, et al.
1556
HMA4 expression in tobacco reduces Cd accumulation due to the induction of the apoplastic
1557
barrier. Journal of Experimental Botany. 2014;65:1125-39.
ip t
1555
Siemianowski O, Mills RF, Williams LE, Antosiewicz DM. Expression of the P-1B-type
1559
ATPase AtHMA4 in tobacco modifies Zn and Cd root to shoot partitioning and metal tolerance.
1560
Plant Biotechnol J. 2011;9:64-74.
us
1562
Sinclair SA, Kramer U. The zinc homeostasis network of land plants. Bba-Mol Cell Res. 2012;1823:1553-67.
an
1561
cr
1558
Song CP, Guo Y, Qiu QS, Lambert G, Galbraith DW, Jagendorf A, et al. A probable
1564
Na+(K+)/H+ exchanger on the chloroplast envelope functions in pH homeostasis and
1565
chloroplast development in Arabidopsis thaliana. Proceedings of the National Academy of
1566
Sciences of the United States of America. 2004;101:10211-16.
d
M
1563
Song WY, Choi KS, Kim DY, Geisler M, Park J, Vincenzetti V, et al. Arabidopsis PCR2 Is a
1568
Zinc Exporter Involved in Both Zinc Extrusion and Long-Distance Zinc Transport. Plant Cell.
1569
2010;22:2237-52.
Ac ce p
1570
te
1567
Stacey MG, Patel A, McClain WE, Mathieu M, Remley M, Rogers EE, et al. The
1571
Arabidopsis AtOPT3 protein functions in metal homeostasis and movement of iron to
1572
developing seeds. Plant Physiology. 2008;146:589-601.
1573
Sunarpi, Horie T, Motoda J, Kubo M, Yang H, Yoda K, et al. Enhanced salt tolerance
1574
mediated by AtHKT1 transporter-induced Na+ unloading from xylem vessels to xylem
1575
parenchyma cells. Plant Journal. 2005;44:928-38.
1576
Suzuki M, Tsukamoto T, Inoue H, Watanabe S, Matsuhashi S, Takahashi M, et al.
1577
Deoxymugineic acid increases Zn translocation in Zn-deficient rice plants. Plant Molecular
1578
Biology. 2008;66:609-17. 65 Page 65 of 78
1579 1580
Swarbreck SM, Colaco R, Davies JM. Plant calcium-permeable channels. Plant Physiology. 2013;163:514-22. Takahashi R, Bashir K, Ishimaru Y, Nishizawa NK, Nakanishi H. The role of heavy-metal
1582
ATPases, HMAs, in zinc and cadmium transport in rice. Plant signaling & behavior.
1583
2012a;7:1605-7.
ip t
1581
Takahashi R, Ishimaru Y, Senoura T, Shimo H, Ishikawa S, Arao T, et al. The OsNRAMP1
1585
iron transporter is involved in Cd accumulation in rice. Journal of Experimental Botany.
1586
2011;62:4843-50.
us
cr
1584
Takahashi R, Ishimaru Y, Shimo H, Ogo Y, Senoura T, Nishizawa NK, et al. The OsHMA2
1588
transporter is involved in root-to-shoot translocation of Zn and Cd in rice. Plant Cell and
1589
Environment. 2012b;35:1948-57.
an
1587
Tanoi K, Saito T, Iwata N, Kobayashi NI, Nakanishi TM. The analysis of magnesium
1591
transport system from external solution to xylem in rice root. Soil Sci Plant Nutr. 2011;57:265-
1592
71.
d
M
1590
Tapken W, Ravet K, Pilon M. Plastocyanin Controls the Stabilization of the Thylakoid
1594
Cu-transporting P-type ATPase PAA2/HMA8 in Response to Low Copper in Arabidopsis. Journal
1595
of Biological Chemistry. 2012;287:18544-50.
Ac ce p
1596
te
1593
Tian SK, Lu LL, Yang XE, Labavitch JM, Huang YY, Brown P. Stem and leaf sequestration
1597
of zinc at the cellular level in the hyperaccumulator Sedum alfredii. New Phytologist.
1598
2009;182:116-26.
1599
Ueno D, Yamaji N, Ma JF. Further characterization of ferric-phytosiderophore
1600
transporters ZmYS1 and HvYS1 in maize and barley. Journal of Experimental Botany.
1601
2009;60:3513-20.
1602
van Kleeff PJM, Jaspert N, Li KW, Rauch S, Oecking C, de Boer AH. Higher order
1603
Arabidopsis 14-3-3 mutants show 14-3-3 involvement in primary root growth both under
1604
control and abiotic stress conditions. Journal of Experimental Botany. 2014;65:5877-88. 66 Page 66 of 78
1605
Vert G, Barberon M, Zelazny E, Seguela M, Briat JF, Curie C. Arabidopsis IRT2
1606
cooperates with the high-affinity iron uptake system to maintain iron homeostasis in root
1607
epidermal cells. Planta. 2009;229:1171-79. Vert G, Grotz N, Dedaldechamp F, Gaymard F, Guerinot ML, Briat JF, et al. IRT1, an
1609
Arabidopsis transporter essential for iron uptake from the soil and for plant growth. Plant Cell.
1610
2002;14:1223-33.
cr
ip t
1608
Very AA, Nieves-Cordones M, Daly M, Khan I, Fizames C, Sentenac H. Molecular biology
1612
of K+ transport across the plant cell membrane: What do we learn from comparison between
1613
plant species? Journal of Plant Physiology. 2014;171:748-69.
us
1611
Voelker C, Gomez-Porras JL, Becker D, Hamamoto S, Uozumi N, Gambale F, et al. Roles
1615
of tandem-pore K plus channels in plants - a puzzle still to be solved. Plant Biology. 2010;12:56-
1616
63.
1618
M
Wang Y, Wu WH. Potassium Transport and Signaling in Higher Plants. Annual Review of Plant Biology. 2013;64:451-76.
d
1617
an
1614
Waters BM, Grusak MA. Whole-plant mineral partitioning throughout the life cycle in
1620
Arabidopsis thaliana ecotypes Columbia, Landsberg erecta, Cape Verde Islands, and the
1621
mutant line ysl1ysl3. New Phytologist. 2008;177:389-405.
Ac ce p
1622
te
1619
White PJ, Bowen HC, Demidchik V, Nichols C, Davies JA. Genes for calcium-permeable
1623
channels in the plasma membrane of plant root cells. Bba-Biomembranes. 2002;1564:299-309.
1624
White PJ, Broadley MR. Calcium in plants. Annals of Botany. 2003;92:487-511.
1625 1626 1627 1628
White PJ, Brown PH. Plant nutrition for sustainable development and global health.
Annals of Botany. 2010;105:1073-80. Wu B, Becker JS. Imaging techniques for elements and element species in plant science. Metallomics. 2012;4:403-16.
67 Page 67 of 78
1629
Xu WF, Jia LG, Baluska F, Ding GC, Shi WM, Ye NH, et al. PIN2 is required for the
1630
adaptation of Arabidopsis roots to alkaline stress by modulating proton secretion. Journal of
1631
Experimental Botany. 2012a;63:6105-14. Xu WF, Jia LG, Shi WM, Baluska F, Kronzucker HJ, Liang JS, et al. The Tomato 14-3-3
1633
Protein TFT4 Modulates H+ Efflux, Basipetal Auxin Transport, and the PKS5-J3 Pathway in the
1634
Root Growth Response to Alkaline Stress. Plant Physiology. 2013a;163:1817-28.
cr
ip t
1632
Xu WF, Jia LG, Shi WM, Liang JS, Zhou F, Li QF, et al. Abscisic acid accumulation
1636
modulates auxin transport in the root tip to enhance proton secretion for maintaining root
1637
growth under moderate water stress. New Phytologist. 2013b;197:139-50.
us
1635
Xu WF, Shi WM, Jia LG, Liang JS, Zhang JH. TFT6 and TFT7, two different members of
1639
tomato 14-3-3 gene family, play distinct roles in plant adaption to low phosphorus stress. Plant
1640
Cell and Environment. 2012b;35:1393-406.
M
an
1638
Yamasaki H, Hayashi M, Fukazawa M, Kobayashi Y, Shikanai T. SQUAMOSA Promoter
1642
Binding Protein-Like7 Is a Central Regulator for Copper Homeostasis in Arabidopsis. Plant Cell.
1643
2009;21:347-61.
te
d
1641
Yang YQ, Qin YX, Xie CG, Zhao FY, Zhao JF, Liu DF, et al. The Arabidopsis Chaperone J3
1645
Regulates the Plasma Membrane H+-ATPase through Interaction with the PKS5 Kinase. Plant
1646
Cell. 2010;22:1313-32.
1647
Ac ce p
1644
Yao X, Horie T, Xue SW, Leung HY, Katsuhara M, Brodsky DE, et al. Differential Sodium
1648
and Potassium Transport Selectivities of the Rice OsHKT2;1 and OsHKT2;2 Transporters in Plant
1649
Cells. Plant Physiology. 2010;152:341-55.
1650
Yao YN, Xu G, Mou DL, Wang JR, Ma JB. Subcellular Mn compartation, anatomic and
1651
biochemical changes of two grape varieties in response to excess manganese. Chemosphere.
1652
2012;89:150-57.
1653 1654
Yokosho K, Yamaji N, Ueno D, Mitani N, Ma JF. OsFRDL1 Is a Citrate Transporter Required for Efficient Translocation of Iron in Rice. Plant Physiology. 2009;149:297-305. 68 Page 68 of 78
1655 1656
Yuan M, Li X, Xiao J, Wang S. Molecular and functional analyses of COPT/Ctr-type copper transporter-like gene family in rice. BMC Plant Biology. 2011;11:69.
1657
plants. Plant and Soil. 2010;326:45-60.
ip t
1658
Zhang JL, Flowers TJ, Wang SM. Mechanisms of sodium uptake by roots of higher
Zhang JL, Flowers TJ, Wang SM. Differentiation of low-affinity Na+ uptake pathways
1660
and kinetics of the effects of K+ on Na+ uptake in the halophyte Suaeda maritima. Plant and
1661
Soil. 2013;368:629-40.
cr
1659
Zhang WJ, To JPC, Cheng CY, Schaller GE, Kieber JJ. Type-A response regulators are
1663
required for proper root apical meristem function through post-transcriptional regulation of
1664
PIN auxin efflux carriers. Plant Journal. 2011;68:1-10.
an
us
1662
Zhang Y, Xu YH, Yi HY, Gong JM. Vacuolar membrane transporters OsVIT1 and OsVIT2
1666
modulate iron translocation between flag leaves and seeds in rice. Plant Journal. 2012;72:400-
1667
10.
M
1665
Zhao HJ, Wu LQ, Chai TY, Zhang YX, Tan JJ, Ma SW. The effects of copper, manganese
1669
and zinc on plant growth and elemental accumulation in the manganese-hyperaccumulator
1670
Phytolacca americana. Journal of Plant Physiology. 2012;169:1243-52.
1672 1673
te
Ac ce p
1671
d
1668
Zhao J, Cheng NH, Motes CM, Blancaflor EB, Moore M, Gonzales N, et al. AtCHX13 is a
plasma membrane K+ transporter. Plant Physiology. 2008;148:796-807. Zheng LQ, Yamaji N, Yokosho K, Ma JF. YSL16 Is a Phloem-Localized Transporter of the
1674
Copper-Nicotianamine Complex That Is Responsible for Copper Distribution in Rice. Plant Cell.
1675
2012;24:3767-82.
1676 1677
Zörb C, Senbayram M, Peiter E. Potassium in agriculture – Status and perspectives. Journal of Plant Physiology. 2014;171:656-69.
1678
Zorrig W, Abdelly C, Berthomieu P. The phylogenetic tree gathering the plant
1679
Zn/Cd/Pb/Co P1B-ATPases appears to be structured according to the botanical families.
1680
Comptes rendus biologies. 2011;334:863-71.
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Table 1. Abbreviations list Ca2+-ATPase
BCD
Bush-and-chlorotic-dwarf
CAX
Ca2+/H+ exchanger
CDF
Cation diffusion facilitator
CEC
Cation exchange capacity
CHX
Cation/H+ exchanger
CNGC
Cyclic-nucleotide gated channel
COPT
Copper transporter
DA-NSCC
Depolarization-activated nonselective cation channel
DMA
Deoxymugineic acid
ECA
Endomembrane-type Ca2+-ATPase
ENA
Efflux transporter of nicotianamine
FRD
Ferric reductase defective
FRDL
Ferric reductase defective-like
FRO
Ferric-chelate reductase oxidase
GLR
Ionotropic glutamate receptor
GSH
Glutathione
HA
H+-ATPase
HA-NSCC
Hyperpolarization-activated nonselective cation channel
HAK/KT/KUP
K+/H+ symporters
HATS
High-affinity transporter system
HKT/Trk
K+/Na+ symporters
HMA
Heavy metal associated
IREG/FPN
Iron-regulated/Ferroportin
IRT
Iron-regulated transporter
KCO
Kir-like K+ channel
KEA
K+ efflux antiporter
LATS
Low-affinity transporter system
LCT
Low-affinity cation transporter
MA
Mugineic acid
MATE
Multidrug and toxic compound extrusion
MRS2/MGT
Magnesium transporter
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MIT
Mitochondrial transporter
MT
Metallothionein
MTP
Metal tolerance proteins
NA
Nicotianamine
NHX
Na+/H+ antiporter
NRAMP
Natural resistance associated macrophage protein
NSCC
Nonselective cation channel
OM
Organic matter
OPT
Oligopeptide transporter
PC
Phytochelatin
PCR
Plant cadmium resistance
PEZ
Phenolics efflux zero
PIC
Permease in chloroplast
PS
Phytosiderophore
RDA
Recommended dietary allowances
TPC
Two-pore channel
TPK
Two-pore K+ channel
TOM
Transporter of mugineic acid family phytosiderophores
VI-NSCC
Voltage-insensitive nonselective cation channel
VIT
Vacuolar iron transporter
YS
Yellow stripe
YSL
Yellow stripe-Like
ZIF
Zinc-induced facilitator
ZIP
(ZRT-IRT)-like protein
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Mg2+/H+ antiporter
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MHX
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S0176-1617(15)00061-9 http://dx.doi.org/doi:10.1016/j.jplph.2015.02.010 JPLPH 52141
To appear in: Received date: Revised date: Accepted date:
15-12-2014 11-2-2015 11-2-2015
Please cite this article as: Pinto E, Ferreira I, CATION TRANSPORTERS/CHANNELS IN PLANTS: TOOLS FOR NUTRIENT BIOFORTIFICATION, Journal of Plant Physiology (2015), http://dx.doi.org/10.1016/j.jplph.2015.02.010 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
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CATION TRANSPORTERS/CHANNELS IN PLANTS:
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TOOLS FOR NUTRIENT BIOFORTIFICATION
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Edgar Pinto1, 2 *, Isabel Ferreira1
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REQUIMTE/ Department of Chemical Sciences, Laboratory of Bromatology and Hydrology, Faculty of Pharmacy - University of Porto, Portugal.
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CISA – Research Centre on Environment and Health, School of Allied Health
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Sciences, Polytechnic Institute of Porto, Portugal.
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Corresponding author address: Rua Jorge Viterbo Ferreira, 228, 4050-313 Porto,
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Portugal; email: [email protected]; Tel: +351220428642; Fax: +351226093390
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Abstract
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Cation transporters/channels are key players in a wide range of physiological functions
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in plants, including cell signaling, osmoregulation, plant nutrition and metal tolerance.
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The recent identification of genes encoding some of these transport systems has allowed
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new studies towards further understanding of their integrated roles in plant. This review
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summarizes recent discoveries regarding the function and regulation of the multiple
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systems involved in cation transport in plant cells. The role of membrane transport in
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the uptake, distribution and accumulation of cations in plant tissues, cell types and
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subcellular compartments is described. We also discuss how the knowledge of inter-
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and intra-species variation in cation uptake, transport and accumulation as well as the
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molecular mechanisms responsible for these processes can be used to increase nutrient
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phytoavailability and nutrients accumulation in the edible tissues of plants. The main
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trends for future research in the field of biofortification are proposed.
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Introduction
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Plant cells contain several cations. Potassium (K), calcium (Ca) and magnesium (Mg)
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are the major cationic species (Maathuis, 2009); iron (Fe), copper (Cu), manganese
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(Mn), nickel (Ni) and zinc (Zn) are widely recognized to be trace essential cations
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(Pilon et al., 2009) and cobalt (Co) and sodium (Na) are known as “beneficial” elements
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(Pilon-Smits et al., 2009).
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Each of the essential cations has a distinct specific role. Both Ca and Mg are
35
constituents of organic structures, such as enzyme molecules, where they may be
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involved in the catalytic function of the enzymes (Maathuis, 2009). Potassium is one of
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the few cations that are not a component of organic structures. It is rather involved in
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the osmoregulation, the maintenance of electrochemical balance in cells and their
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compartments, and the regulation of the enzymatic activity (Wang and Wu, 2013). Due
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to their low concentration, trace essential cations do not play a direct role in
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osmoregulation or similar processes. These elements are redox-active and act as
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catalytically active cofactors in enzymes. Others have enzyme-activating functions, and
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others fulfil a structural role in stabilizing proteins (Hänsch and Mendel, 2009, Pilon-
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Smits et al., 2009).
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Cations must be acquired by plant roots from the soil solution and then delivered to
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tissues, organs and subcellular compartments, where they are required to support plants’
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metabolism (Maathuis, 2009). Plants face major variations in cation uptake, particularly
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in the case of trace essential (Fe, Zn, Cu, Ni and Mn) and beneficial (Co and Na)
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cations. On the one hand, the phytoavailability of these elements can be low, resulting
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in deficiency symptoms of the plants that grown in such environments. On the other
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hand, toxicity can occur in certain environments were the phytoavailability of
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problematic elements (e.g., Na) is enhanced by the conditions of that environment
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(Pinto et al., 2014b, Pinto et al., 2015). Plants have to cope with variable demands of
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different tissues during growth or in response to abiotic stresses. Thus, along the course
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of evolution, plants have developed highly specialized homeostatic mechanisms that
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deal with systemic spatiotemporal requirements in the acquisition, transport and supply
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of cations to specific tissues, organs or compartments (Pinto et al., 2014a).
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It is been widely recognized that the distinct patterns in the uptake, distribution and
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accumulation of cations in plants is the result of selective transport processes in which
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membrane transport proteins are involved (Haferkamp and Linka, 2012). The
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identification of several gene families encoding putative cation transporters/channels
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constituted a major breakthrough in the plant science field enabling the realization of
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studies at the gene and protein levels (e.g., gene expression at the tissue or plant levels,
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transport activity and subcellular localization) to achieve a further understanding of the
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integrated roles of these transport systems in plants.
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The list of essential minerals for plants is quite similar but not equal to the list for
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humans. For example, Se, I, Co and Na are essential for humans but not for most plants
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(White and Brown, 2010). Therefore, human deficiencies may arise for these elements
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in populations that depend on plants grown locally where the soil content of these
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elements is low. Actually, serious human deficiencies of Se and I exist worldwide
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(Hirschi, 2009). For nutrients required by both plants and humans, one can expect that
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human deficiencies will be less of a problem because the elements will necessarily be
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present in plant foods. However, the mineral content of plants is sometimes very low to
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meet human requirements, or the minerals may be present in forms that cannot be
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efficiently utilized by humans. Examples of both these situations are Ca and Fe,
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respectively (Jeong and Guerinot, 2008, Dayod et al., 2010). Thus, new strategies are
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needed to ensure a sustainable supply of healthy food to world’s population while
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meeting human nutritional requirements.
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The aim of this review is to join together information about the major cation
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transporters/channels involved in the uptake, distribution and accumulation of essential
81
(K, Ca, Mg, Fe, Cu, Mn, Zn and Ni) and beneficial (Co and Na) elements and how this
82
information can be used in biofortification programs. In the following sections, a
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detailed description of the transporters/channels function will be provided taking into
84
consideration the four major processes that elements usually undergo – (1) uptake, (2)
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root compartmentation, (3) translocation and (4) leaf compartmentation. To simplify, in
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each of these four processes the transporters/channels involved in cation transport will
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be described according to the following division: (i) macroelements (K, Ca and Mg);
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(ii) trace essential elements (Fe, Cu, Mn, Ni and Zn); and (iii) beneficial elements (Co
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and Na). Then, a careful description of pathways to improve the biofortification of plant
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foods will be provided.
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Uptake
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Cation transport across the plant plasma membrane is mediated by an electrochemical
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gradient of protons generated by plasma membrane H+-ATPases. This “primary
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transport system” pumps protons out of the cell creating pH and electrical potential
96
differences across the plasma membrane (Fox and Guerinot, 1998). At that point,
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“secondary transport systems” can utilize these gradients for many plant functions such
98
as nutrients influx and efflux (Palmgren, 2001b). Cations enter the root either by
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crossing the plasma membrane of the root endodermal cells (symplastic transport) or by
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entering the root apoplast through the space between cells (apoplastic transport). They
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cannot pass through membranes without the aid of membrane transporter proteins.
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These transporter proteins occur naturally in several plant membranes (e.g., tonoplast,
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endoplasmic reticulum, mitochondria, chloroplasts) and, usually, multiple transporters
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exist for each element (Pinto et al., 2014a). For instance, rice (Oryza sativa) has at least
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seven Cu transporters (Yuan et al. , 2011). Each transporter has unique properties.
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When a low concentration of nutrients is present in the soil solution, their uptake
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usually requires the high-affinity transport system (HATS). By contrast, the low-affinity
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transport system (LATS) is used more when high concentrations of nutrients are
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present, such as in agricultural soils after fertilization. Furthermore, the abundance of
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each transporter varies with tissue type and environmental conditions, making the
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uptake of cations in plants a complex process (Karley and White, 2009, Puig and
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Penarrubia, 2009).
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Macroelements
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Potassium (K) is the most abundant cation in plant cells and plays a vital role in plant
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growth and development. Normally, K content in soil solution ranges between 1 and 10
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mM whereas intracellular K levels are maintained at 100 – 200 mM. Therefore, K
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uptake from the soil is obviously against a concentration gradient (Anschutz et al.,
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2014, Zörb et al., 2014). Similarly to most mineral nutrients, the main acquisition of K
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from the outside environment follows a biphasic form, defined as the sum of two
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distinguishable uptake mechanisms (HATS and LATS) at the root plasma membrane
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(Wang and Wu, 2013). Several genes appear to encode putative K transporters/channels
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in the plasma membrane of Arabidopsis root cells (Figure 1). They are grouped into
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four families, including HAK/KT/KUP (K+/H+ symporters), HKT/Trk (K+/Na+
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symporters), CHX (cation/H+ antiporters) and Shaker channels (Nieves-Cordones et al.,
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2014). The bulk of HATS that mediate the influx of K are catalyzed by members of the
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HAK/KT/KUP family (Very et al., 2014). In Arabidopsis, 10 of 13 AtKT/KUPs are
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expressed in root hairs. Five of them were expressed in root tip cells suggesting an
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essential role of these HATS in K uptake (Ahn et al., 2004). From these 5 AtKT/KUPs
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expressed in Arabidopsis roots, AtKUP1, AtKUP4 and AtHAK5 are transporters in the
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plasma membrane of root cells that seem to be engaged in the K influx (Ahn et al.,
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2004, Fulgenzi et al., 2008, Rubio et al., 2008). The physiological importance of
133
HKT/Trk transporters for high-affinity K influx is not yet fully clear. Several plant
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studies have pointed out that most of HKT transporters function as Na transporters and
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only a limited number of these transporters are Na+/K+ symporters. Therefore, evidence
136
supports a restricted role of the HKT family in K uptake (Very el al., 2014). Several
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members of the CHX family have been identified as K transporters. For example,
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AtCHX13 and AtCHX17 have been proposed as possible pathways for root K influx
139
(Cellier et al., 2004, Zhao et al., 2008). Low-affinity K influx seems to be mediated by
140
Shaker K-channels such as AtAKT1 and AtKC1, which are mainly expressed in root
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epidermal cells (Geiger et al., 2009, Ardie et al., 2010, Jeanguenin et al., 2011). Plant
142
Shaker channels display extraordinary K selectivity. Consequently, the Shaker K-
143
channels AtAKT1 and AtKC1 as well as the HAK/KT/KUP transporter AtHAK5 are
144
responsible for almost all K influx (Nieves-Cordones et al., 2014).
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A peculiar electrochemical gradient exists for Ca in plant roots since cytosolic Ca levels
146
are in the sub-µM range, while Ca concentrations in the soil solution are in the mM
147
range (Karley and White, 2009). Calcium is taken up symplastically through
148
nonselective cation channels (NSCCs) present in the root system. These can be divided
149
according to their voltage dependence into depolarization-activated (DA-NSCCs),
150
hyperpolarization-activated (HA-NSCCs) and voltage-insensitive (VI-NSCCs) types, as
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shown in Figure 1 (Demidchik and Maathuis, 2007). Regarding HA-NSCCs, members
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of the annexin gene family are likely to encode these Ca channels (Mortimer et al.,
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2008). For VI-NSCCs, two gene families, the plasma membrane-localized cyclic-
154
nucleotide gated channels (CNGC) and the ionotropic glutamate receptor (GLR)
155
homologues are proposed to mediate Ca uptake (Ma et al., 2009, Swarbreck et al., 2013,
156
Forde, 2014). Despite the type of NSCC used to mediate Ca influx, the opening of these
157
channels must be tightly regulated, because changes in cytosolic Ca levels trigger
158
several specific responses to many environmental and developmental stimuli (McAinsh
159
and Pittman, 2009).
160
Magnesium concentration in soil solutions normally ranges from 8.5 to 125 mM.
161
Cytosolic Mg levels are around 0.4 mM, thus Mg influx into root cells can be achieved
162
by Mg-permeable cation channels (Karley and White, 2009). Despite that, members of
163
the MRS2/MTG gene family expressed in Arabidopsis appear to dominate the Mg
164
uptake across the root plasma membrane (Figure 1) (Gebert et al., 2009). A member of
165
the A. thaliana Mg transport family, AtMGT1, was expressed in tobacco (Nicotiana
166
benthamiana) and showed an increased Mg content (Deng et al., 2006). In addition to
167
AtMGT1, other two MRS2/MTG members, AtMGT7 and AtMGT9, are expressed in
168
Arabidopsis roots and have a key role in Mg uptake (Mao et al., 2008, Chen et al.,
169
2009).
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Trace essential elements
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Several Fe species may be taken up by plant roots. The divalent form (Fe2+) is the main
173
absorbed species but Fe3+ and Fe-chelates are also taken up. Plants have developed
174
several mechanisms to enhance Fe uptake. Two main categories can be distinguished
175
according to the plant species: Strategy I in non-graminaceous plants and Strategy II in
176
graminaceous plants (Figure 2). The main processes of Strategy I (used by non-
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graminaceous plants such as A. thaliana) are the excretion of protons and phenolic
178
compounds from the roots to the rhizosphere, reduction of ferric chelates at the root
179
surface, and the absorption of the resulting ferrous ions across the root by plasma
180
membrane transporters (Kobayashi and Nishizawa, 2012). The H+-ATPase (HA)
181
transporters are responsible for the efflux of protons to promote the acidification of the
182
rhizosphere (Santi and Schmidt, 2009) and the recently identified OsPEZ1 and OsPEZ2
183
(phenolics efflux zero) seems to be involved in the secretion of phenolics (Bashir et al.,
184
2011a, Ishimaru et al., 2011). Non-graminaceous plants depend on ferric reductase
185
oxidase (e.g., AtFRO2) to reduce the ferric chelates at the root surface (Jeong and
186
Connolly, 2009) and then on the iron-regulated transporter (AtIRT1, AtIRT2 and
187
AtIRT3) to absorb the generated Fe2+ ions across the root plasma membrane (Lin et al.,
188
2009, Vert et al., 2009, Barberon et al., 2011). The Strategy II, used by graminaceous
189
plants (e.g., barley, maize and rice), depends on the biosynthesis and secretion of
190
phytosiderophores (PS) such as mugineic acids (MAs) which mobilize Fe (Kobayashi
191
and Nishizawa, 2012). Recently, the gene responsible for MA secretion was identified
192
to be the transporter of the mugineic acid family phytosiderophores 1 (OsTOM1 and
193
HvTOM1, for rice and barley, respectively) (Nozoye et al., 2011). The MAs secreted
194
into the rhizosphere stimulate the solubilization of Fe3+ in the soil and the resulting Fe-
195
PS complexes can be taken up through Yellow Sripe (YS) and Yellow Stripe-like (YSL)
196
transporters such as HvYS1, ZmYS1 and OsYSL15 (Murata et al., 2008, Inoue et al.,
197
2009, Ueno et al., 2009, Nozoye et al., 2013). Despite being Strategy II plants,
198
graminaceous plants (e.g., rice), can also have ferrous transporters (e.g., OsIRT1,
199
OsNRAMP1 and OsNRAMP5) allowing the absorption of Fe2+ in addition to its
200
Strategy II–based Fe3+-MA uptake (Takahashi et al., 2011, Ishimaru et al., 2012).
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The most bioavailable form of Cu in soils is Cu2+. However, root uptake is often
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facilitated by reduction of Cu2+ to Cu+. The ferric reductase oxidases AtFRO2, AtFRO3,
203
AtFRO4 and AtFRO5, expressed in A. thaliana, seem to be involved in this process
204
(Mukherjee et al., 2006, Bernal et al., 2012). Copper is likely to enter the cytosol of root
205
cells through a cell surface COPT/Ctr-family transporter. Six members of the Ctr family
206
(COPT1-6) which mediate the influx of Cu have been identified in Arabidopsis.
207
AtCOPT1, AtCOPT2 and AtCOPT3 are expressed in the plasma membrane of roots and
208
have an important role in the acquisition of Cu monovalent ions (Figure 3) (Yamasaki et
209
al., 2009, Andres-Colas et al., 2010). Furthermore, members of the Zinc-Regulated
210
Transporter and Iron-Regulated Transporter (ZRT-IRT)-like proteins (ZIP) family can
211
also mediate the uptake and transport of Cu in Arabidopsis (Yamasaki et al., 2009, del
212
Pozo et al., 2010). Members of the YS and YSL family (e.g., ZmYS1 and HvYSL2) are
213
also involved in the transport of Cu-PS complexes (Murata et al., 2008, Araki et al.,
214
2011).
215
The oxidized forms Mn3+ and Mn4+ are not bioavailable to plants. It is the reduced form
216
of this element, Mn2+, that is absorbed by root cells (Pittman, 2005). The plasma
217
membrane IRT1, expressed in both Arabidopsis and barley, can transport Mn (Figure 3)
218
(Pedas et al., 2008, Barberon et al., 2011). The overexpression of AtIRT2 leads to
219
overaccumulation of Mn in transgenic plants, but the role of this transporter in Mn
220
uptake remains to be clarified (Vert et al., 2009). The plasma membrane-localized
221
NRAMP1 and NRAMP5 were shown to be high-affinity Mn transporters in Arabidopsis
222
and rice, respectively. Both AtNRAMP1 and OsNRAMP5 have broad selectivity, and
223
their expression is restricted to the root (Cailliatte et al., 2010, Ishimaru et al., 2012). In
224
maize and barley, Mn uptake can also be performed by the ZmYS1 and HvYSL2
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transporters, respectively, which were confirmed to be capable of acquiring Mn-PS
226
complexes from the rhizosphere (Murata et al., 2008, Araki et al., 2011).
227
Nickel is present in the environment usually in the form of Ni2+, which is the most
228
available form to plants. Divalent cations such as Ni, Cu and Fe have very similar
229
physicochemical properties thus they compete with each other in biochemical and
230
physiological processes in plants (Ghasemi et al., 2009). Nickel can enter symplastically
231
into the root cells through AtIRT1 (Nishida et al., 2012). Moreover, Ni can also be
232
taken up in the form of Ni-PS complexes by the ZmYS1 due to its broad selectivity for
233
several cations (Figure 3) (Murata et al., 2008).
234
Zinc is taken up across the plasma membrane of root cells as a free ion and/or
235
complexed with PS (Sinclair and Kramer, 2012). The ZIP transporter family includes
236
the best candidates for facilitating Zn influx into the plant cytoplasm. In addition to its
237
role in Fe acquisition, the broad selectivity of AtIRT1 allows this transporter to mediate
238
also the uptake of several divalent metal cations, including Zn (Figure 3) (Barberon et
239
al., 2011, Fukao et al., 2011, Shanmugam et al., 2011). This was also demonstrated in
240
rice plants overexpressing OsIRT1, which accumulate elevated levels of Zn in the
241
shoots, roots and mature seeds (Lee and An, 2009). Like their close homologue AtIRT1,
242
AtIRT2 and AtIRT3 are also able to transport Zn into roots (Lin et al., 2009, Vert et al.,
243
2009). Other ZIP transporters are involved in the uptake of Zn from soil. For example,
244
OsZIP1 and OsZIP3 are likely to play a role in this process (Bashir et al., 2012). In
245
Arabidopsis, AtZIP4 expression is induced upon Zn deficiency, showing its function as
246
a Zn transporter (Assunção et al., 2010). The rice plasma membrane-localized OsZIP8
247
also seems to be a Zn transporter that functions in Zn uptake (Lee et al., 2010). A
248
member of the NRAMP family, AtNRAMP1, is also able to mediate Zn influx into the
249
root cells of Arabidopsis (Cailliatte et al., 2010). Zinc is also taken up from the
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250
rhizosphere in the form of Zn–PS complexes by members of the YS and YSL family
251
(Murata et al., 2008, Araki et al., 2011).
252
Beneficial elements
254
Since soil salinity is a global environmental challenge and NaCl is typically the major
255
salt that contributes to that salinity, much research activity has been dedicated to the
256
characterization of Na transport and distribution in plants, particularly to its uptake by
257
plant roots (Zhang et al., 2010). Sodium can enter the root cells by NSCCs, more
258
specifically by the VI-NSCCs. In fact, VI-NSCC permeability to Na has been
259
demonstrated in several tissues and species, resulting in substantial evidence that root
260
Na influx is mediated by these channels (Demidchik and Maathuis, 2007). Another
261
candidate for mediating Na influx is the low-affinity cation transporter (LCT) 1.
262
However, this function is still not clear and further investigation must be carried out
263
(Amtmann et al., 2001). K and Na ions compete with each other to enter plant cells due
264
to their physicochemical similarity. Therefore, it is plausible that K transporters may
265
also be involved in Na acquisition and distribution. However, the involvement of K
266
transporters from both the HAK/KT/KUP and AKT families in Na movement is
267
somewhat contradictory. Some studies support the idea that some members of the
268
HAK/KT/KUP and AKT families are involved in Na accumulation (Ardie et al., 2010,
269
Horie et al., 2011, Zhang et al., 2013) but others fail to confer a specific role to these
270
transporters in Na influx (Nieves-Cordones et al., 2010). Significant evidence exists for
271
the involvement of K transporters from the HKT families in the Na influx (Benito et al.,
272
2014). The HKT family can be divided in two different sub-families according to their
273
properties regarding Na and K transport in heterologous expression systems. Sub-family
274
1, which shows Na-specific transport activity and mediates Na uptake, includes, among
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others, AtHKT1;1 and its orthologues OsHKT1;1 and HvHKT1;1 expressed in rice and
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barley, respectively (Haro et al., 2005, Sunarpi et al., 2005, Davenport et al., 2007,
277
Jabnoune et al., 2009). Sub-family 2 functions as a K-Na co-transporter and includes,
278
among others, HvHKT2;1, OsHKT2;1, OsHKT2;2 and OsHKT2;4 (Yao et al., 2010,
279
Mian et al., 2011, Sassi et al., 2012). It should be noted that the involvement of a certain
280
transporter may be genotype-specific and not transversal to all plant species.
281
The mechanisms of Co uptake remain poorly understood, although some transporters
282
have been identified. Cobalt can be taken up in its ionic form Co2+ through the plasma
283
membrane-localized AtIRT1 (Vert et al., 2002) and in the form of Co-PS complex by
284
the ZmYS1 in maize (Murata et al., 2008). However, other membrane transporter,
285
AtNRAMP1, shows broad selectivity for divalent cations and can also participate in the
286
influx of Co into root cells (Cailliatte et al., 2010).
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Root compartmentation
289
Once inside plant root cells, cations can be stored in vacuoles, transformed and/or
290
translocated to shoot tissues. Evidence suggests that monovalent cations have a crucial
291
role in the osmotic potential of cells. By contrast, divalent cations, like Ca 2+, are usually
292
complexed with organic compounds within cells and thus, are responsible for other
293
tasks inside plant tissues (Martinoia et al., 2012). Plants have developed effective
294
mechanisms to perform the chelation and sequestration of cations, such as induction of
295
higher levels of chelating species, compartmentation in vacuoles and adsorption to the
296
cell wall (Figure 4). Nicotianamine (NA), glutathione (GSH), phytochelatins (PCs),
297
metallothioneins (MTs), and organic acids have been increasingly shown in the
298
literature to play a key role in the chelation and sequestration of several cations (Guo et
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al., 2008, Larbi et al., 2010, Pal and Rai, 2010, Deinlein et al., 2012, Haydon et al.,
300
2012).
301
With respect to root compartmentation, cations may be stored in the vacuole and/or cell
302
wall. The vacuole is generally considered to be the main storage site for cations in plant
303
cells and vacuolar compartmentation of elements is also a part of the tolerance
304
mechanism (Martinoia et al., 2012). However, cell wall proteins and pectins are also
305
involved in metal chelation and tolerance (Larras et al., 2013).
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Macroelements
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In addition to its role in the maintenance of cell osmoticum, K is also required in several
309
biochemical reactions. Since K is less chaotropic than Na, K is preferentially
310
accumulated in plant cells (Wang and Wu, 2013). Once in the cytosol, K can cross the
311
tonoplast through transporters or channels. Despite the limited knowledge of the
312
mechanisms that regulate K movement across the tonoplast, evidence suggests that
313
transporters are responsible for the influx of K into vacuoles while channels mainly
314
mediate its efflux to the cytosol (Martinoia et al., 2012, Ahmad and Maathuis, 2014).
315
The Na+/H+ antiporters (NHX) are a class of transporters that catalyzes the
316
electroneutral exchange of Na and/or K for H+ by using the electrochemical H+ gradient
317
produced by the H+–ATPases present in the plasma membrane of cell organelles
318
(Ahmad and Maathuis, 2014, Pottosin and Dobrovinskaya, 2014). In Arabidopsis
319
thaliana, six intracellular NHX members are identified and divided into two groups,
320
vacuolar (NHX1 to NHX4) and endosomal (NHX5 and NHX6) (Bassil et al., 2011).
321
From the four vacuolar NHX transporters, two (NHX1 and NHX2) have been fully
322
characterized and have a crucial role in the regulation of intra-vacuolar K in root cells
323
(Barragan et al., 2012). Regarding K efflux to the cytosol, two families of K-permeable
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channels, TPC1 (two-pore channel1) and the TPK/KCOs (two-pore K channels), have
325
been identified in the tonoplast of root cells (Martinoia et al., 2012). The first class of K
326
channels is the TPC1 which was the first vacuolar channel to be identified and mediates
327
the Ca-dependent K, Na and Mg efflux from the vacuole (Hedrich and Marten, 2011). It
328
has been suggested a possible role of TPC1 in Ca efflux from the vacuoles but some
329
contradictory data about this issue exist in the literature (Ranf et al., 2008). The
330
TPK/KCO family constitutes the second class of K channels located in the vacuolar
331
membrane. In Arabidopsis thaliana, the AtTPK/KCO family comprises five two-pore K
332
channels (TPKs) and one Kir-type K channels (KCO). From the five TPKs, four
333
(AtTPK1, -2, -3, and -5) are localized in the tonoplast as well as the AtKCO3 (Voelker
334
et al., 2010). Characterization of Arabidopsis tpk1 knockout mutants and TPK1-
335
overexpressing lines demonstrate that TPK1 has an essential role in intracellular K
336
homeostasis by controlling the K vacuolar release (Hamamoto and Uozumi, 2014).
337
Calcium is usually present at high concentrations in the cytosol of root cells, where it
338
has a pivotal role in the stabilization of cell walls, and also in the vacuole, where its
339
content is regulated by a larger number of Ca transporters (Pittman, 2011). The
340
cytosolic concentration of Ca is around 100 nM while the vacuolar concentration is in
341
the mM range. Hence, vacuolar compartmentation of Ca is an energy-dependent process
342
mediated by two classes of Ca vacuolar transporters (Martinoia et al., 2012). The first
343
class includes the P-type Ca-ATPases and includes two Ca transporters, ACA4 and
344
ACA11, localized in the vacuolar membrane. ACA4 occurs mainly on small vacuoles,
345
whereas ACA11 is localized in the large vacuoles of root cells (Lee et al., 2007). The
346
second class of vacuolar Ca transporters includes the Ca2+/H+ exchangers (CAXs). Six
347
CAX transporters have been identified in Arabidopsis thaliana that are split into two
348
phylogenetic groups that seems to reflect their functional variation (Manohar et al.,
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2011). The Type-1B group, which includes CAX2, CAX5 and CAX6, seem to have a
350
broader substrate specificity confirmed by its ability to transport several divalent cations
351
(e.g., Mn and Cd) besides Ca (Edmond et al., 2009). The Type 1A group, including the
352
CAX1, CAX3 and CAX4 transporters, appears to have a higher specificity for Ca
353
transport (Cheng et al., 2005). Arabidopsis cax1 knockout mutants do not suffer
354
significant changes in their Ca content, however, cax1cax3 double mutants exhibited
355
reduced Ca concentration in their vacuoles compared to wild-type plants (Conn et al.,
356
2011b, Connorton et al., 2012).
357
Plants growing on Mg-rich soils use this divalent cation as an osmotic regulator. The
358
MGTs (Mg transporters) are the major transporters of this cation described in plants. In
359
Arabidopsis thaliana, ten members with Mg2+/H+ antiport activity are identified
360
(Martinoia et al., 2012). Two of them, AtMGT2/AtMRS2-1 and AtMGT3/AtMRS2-5,
361
are localized in the vacuolar membrane. Plant lines overexpressing these transporters
362
have shown Mg accumulation (Conn et al., 2011a). Arabidopsis Mg2+/H+ exchanger
363
(AtMHX) is another putative vacuolar Mg2+/H+ antiporter that seems to mediate the Mg
364
influx across the tonoplast (Shaul et al., 1999). The same mechanism was also observed
365
in the hyperaccumulator Arabidopsis halleri expressing an AtMHX homolog (Elbaz et
366
al., 2006).
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368
Trace essential elements
369
Plants require certain micronutrients for proper growth and development. For example,
370
Fe is involved in numerous redox reactions, Cu has a crucial role in electron transfer
371
between photosystem I and II, Mn is required as a cofactor for a variety of enzymes,
372
such as the Mn-dependent superoxide dismutase (MnSOD), and Zn acts as a catalytic or
373
structural co-factor in a large number of enzymes and regulatory proteins (Hänsch and
16 Page 16 of 78
Mendel, 2009). Micronutrients deficiency is a more common reality that toxicity.
375
Hence, plants usually accumulate extra nutrients to sustain their growth in times of
376
need. To accomplish that, micronutrients are translocated to seeds where they can
377
support the growth of new plantlet. Therefore, most of these micronutrients are not
378
retained in plant roots, so this topic is less relevant to discuss. Only a brief overview
379
will be made here.
380
The bulk of Fe is found in leaves chloroplasts, where this element is engaged in several
381
biochemical reactions (Hänsch and Mendel, 2009). A certain amount of this
382
micronutrient can also be stored in the leaves vacuoles or mitochondria (Martinoia et
383
al., 2012). Iron can also be stored in seeds vacuoles (Kim et al., 2006b). Its storage in
384
root cells is not very common.
385
Non-tolerant plant species, such as A. thaliana, can use chelation as a mechanism to
386
avoid Cu-induced damage in the root cells. In a Cu-rich environment, MTs and PCs are
387
usually up-regulated in response to Cu stress (Guo et al., 2008, Pal and Rai, 2010). Four
388
Arabidopsis MTs (AtMT1a, AtMT2a, AtMT2b, and AtMT3) have been implicated in
389
the Cu accumulation and tolerance in roots under high Cu levels (Guo et al., 2008). In
390
addition to the MTs tolerance mechanism, Cu can be sequestered in the vacuole. A
391
member of the HMA (Heavy Metal Associated) family, AtHMA5, mediates the
392
vacuolar Cu influx in Arabidopsis root cells (Andres-Colas et al., 2006). In A. thaliana,
393
one of the six COPTs identified is targeted to the tonoplast. An Arabidopsis copt5
394
knockout mutant exhibits severe defects in vegetative growth and root elongation,
395
accumulating more Cu in the roots and translocating less Cu to the shoots. These results
396
support the fact that AtCOPT5 would act as a vacuolar Cu exporter (Garcia-Molina et
397
al., 2011, Klaumann et al., 2011).
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A large pool of Mn (around 80%) is usually observed in leaf vacuoles. Moreover, a
399
significant accumulation of Mn is also present in leaf chloroplasts where this element is
400
required as the catalytic center for light-induced water oxidation in photosystem II
401
(Pittman, 2005). The previously mentioned AtCAX2 and AtCAX4 transporters that are
402
engaged in the transport of Ca have also the ability to transport Mn into the vacuole
403
(Edmond et al., 2009, Connorton et al., 2012). AtMTP11, which belongs to the cation
404
diffusion facilitator (CDF) family, is implicated in the pre-vacuolar compartmentation
405
of Mn in root cells (Delhaize et al., 2007). In Arabidopsis, the export of Mn from
406
vacuoles can be performed by AtNRAMP3 and AtNRAMP4 which were shown to have
407
a crucial role in the Mn homeostasis within plant tissues (Lanquar et al., 2010). The P-
408
type ATPase AtECA3, mainly expressed in root tissues, participates in the influx of Mn
409
into the Golgi complex (Mills et al., 2008).
410
After uptake into the root symplast, Ni detoxification at the cellular level is achieved by
411
transport of Ni, as a free ion or chelated with appropriate ligands, to the storage sites
412
such as the vacuole and cell wall (Bhatia et al., 2005). A recent study showed that the
413
main mechanism that confers Ni tolerance seems to be vacuolar Ni sequestration (Saito
414
et al., 2010). A member of the IREG/FPN (Iron-Regulated/Ferroportin) family,
415
AtIREG2/FPN2, has been identified in the tonoplast of root cells and mediates vacuolar
416
Ni loading (Schaaf et al., 2006).
417
Once in the root symplast, Zn can be immobilized in the root through transport into
418
vacuoles. Evidence suggests that the speciation of Zn in the root symplast has a direct
419
influence in the extent of Zn immobilization in root vacuoles (Sinclair and Kramer,
420
2012). The vacuolar membrane transporter AtMTP1 has an essential role in
421
detoxification of excessive Zn. When grown in a Zn-rich medium, the mutant line of
422
Arabidopsis that lacks MTP1 was not able to accumulate Zn in vacuoles, unlike wild-
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type roots (Kawachi et al., 2009). The tonoplast-localized AtMTP3 also contributes to
424
basic cellular Zn tolerance and controls Zn partitioning. Overexpression of MTP3
425
increases Zn accumulation in roots of A. thaliana (Arrivault et al., 2006). The P1B-type
426
ATPase subfamily HMA plays a key role in the process of metal allocation or
427
detoxification in plants. At least eight (HMA1-HMA8) members are identified in
428
Arabidopsis thaliana. AtHMA1 to AtHMA4 belong to the group implicated in divalent
429
cation transport whereas AtHMA5 to AtHMA8 act on monovalent Cu ion transport
430
(Zorrig et al., 2011). From the four HMAs involved in divalent cations transport, one of
431
them, AtHMA3, seems to play a role in the detoxification of Zn by participating in its
432
vacuolar sequestration in root cells (Morel et al., 2009). It has been speculated that ZIF1
433
(for Zinc-Induced Facilitator1) is involved in a mechanism of vacuolar Zn sequestration.
434
Specifically, AtZIF1 has been implicated in the transport of Zn complexes (mainly with
435
NA) into the vacuole, since overexpression of ZIF1 leads to strongly enhanced retention
436
of Zn in root vacuoles (Haydon et al., 2012). A member of the ZIP family, AtZIP1, may
437
contribute to the remobilization of Zn from the vacuole to the cytoplasm in Arabidopsis
438
roots (Milner et al., 2013). Moreover, the tonoplast-localized NRAMP4 also seems to
439
mediate the vacuolar Zn export in root cells (Oomen et al., 2009).
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441
Beneficial elements
442
Sodium is a ubiquitous cation present in all soils. Soils can contain high concentrations
443
of Na, which can be readily absorbed by most plants. Since high Na concentrations
444
within the cytosol can be toxic, vacuolar Na storage is essential for plants to survive
445
under high salinity conditions (Martinoia et al., 2012, Benito et al., 2014). As already
446
described, the NHX family is a Na+/H+ antiporter. Several member of the NHX family
447
(NHX1-4) are targeted in the vacuole and are known to be involved in salt tolerance.
19 Page 19 of 78
Furthermore, the endosomal-localized NHX5 and NHX6 also seem to be engaged in
449
this process, suggesting a complex interactive coordination between different cell
450
organelles in response to salinity (Krebs et al., 2010, Bassil et al., 2011).
451
As regard to Co, few studies have addressed its compartmentation. In fact, the only
452
evidence about vacuolar compartmentation of Co comes from the study of Morrissey et
453
al. (2009) who found that AtIREG2/FPN2, besides being involved in the vacuolar
454
sequestration of Ni, also seems to mediate Co transport across the tonoplast in root
455
cells.
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an
456
Translocation
458
Translocation in plants involves a series of steps, as follows: (1) radial transport across
459
the root tissues, which include symplastic transport to pass through the Casparian strip
460
(a waxy coating that prevents solutes from entering the root xylem directly from the soil
461
solution or root apoplast); (2) xylem loading, transport and unloading; (3) xylem-to-
462
phloem transfer; (4) phloem loading, transport and unloading; (5) symplastic movement
463
toward the site of demand; and (6) retranslocation from source or senescing tissue
464
(Kobayashi and Nishizawa, 2012).
465
Although some transporters are highly specialized in the movement of a certain cation,
466
others show low substrate specificity and can translocate different elements (Zorrig,
467
Abdelly, 2011). Some cations are chelated by organic acids, which are involved in metal
468
absorption by plant roots, translocation in the xylem and storage in the leaf cells
469
vacuoles (Karley and White, 2009, Larbi et al., 2010). Cations can also be bound by NA
470
and mobilized by the YSL family transporters (Aoyama et al., 2009, Ishimaru et al.,
471
2010, Harris et al., 2012). It should be emphasized that cation translocation is a complex
472
process involving multiple networks. Specialized proteins can transport elements with
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473
similar characteristics (e.g., oxidative state), inducing competition that can results in
474
deficiency, toxicity and/or accumulation in the above-ground plant tissues (Puig and
475
Penarrubia, 2009).
ip t
476
Macroelements
478
A large fraction of the K taken up by plant roots is translocated to the shoot. Potassium
479
is transported symplastically to the xylem, where the Shaker-like channel AtSKOR,
480
expressed in the root pericycle and stelar parenchyma of Arabidopsis, is thought to be
481
involved in K loading of the xylem (Johansson et al., 2006). The loading of K within the
482
shoot is mainly determined by transpirational water flows and the shoot apoplastic K
483
concentration (Ahmad and Maathuis, 2014). In addition to the transport of K into
484
above-ground tissues, a large shoot-to-root K flux through the phloem is maintained. In
485
Arabidopsis, the redistribution of K from leaves to phloem-fed tissues seems to be
486
mediated by the Shaker-like channel AtAKT2/AKT3, which is mainly expressed in the
487
phloem and xylem parenchyma and has been indicated to be involved in phloem loading
488
and unloading (Gajdanowicz et al., 2011).
489
Calcium is relatively immobile within plant tissues and thus, it is likely that most of the
490
Ca is sequestered in the root cells vacuoles (Maathuis, 2009). Until now, no transporters
491
have been identified to be responsible for Ca xylem loading, although it is possible for
492
Ca to be symplastically transported across the root to the xylem (Karley and White,
493
2009). Evidence suggests that Ca reaches the xylem apoplastically through regions of
494
the root where the Casparian strips are absent or via the cytosol of unsuberised
495
endodermal cells where Casparian strips are present (Baxter et al., 2009). This
496
assumptions support the observation that Ca mobilization to the shoot occurs mainly
497
from the root apex and/or regions of lateral root initiation, where the Casparian strip is
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disrupted (Shaul, 2002, White et al., 2002). Once in the vascular system, Ca mobility is
499
also low. Calcium is transported in the xylem as free ions (Ca2+) or complexed with
500
organic acids (Karley and White, 2009). Additionally, it seems that the rhizosphere Ca
501
concentration has an important effect on the Ca concentration in the xylem sap and
502
transpirational water flows governs Ca delivery within the shoot (White et al., 2002).
503
Magnesium is loaded into the xylem sap, where it is present as free ions (Mg2+) or
504
complexed with organic acids (Karley and White, 2009). To date, little information is
505
available regarding membrane transporters engaged in the Mg xylem loading. Recently,
506
it was suggested that members of the MRS2/MGT family could be involved in the Mg
507
transport system (Tanoi et al., 2011), but the exact transporter has not been identified
508
yet. As for Ca, evidence suggests that, besides the symplastic pathway, Mg can be
509
loaded into the xylem vessels apoplastically and it seems that this pathways is the
510
preferred for xylem Mg loading (Shaul, 2002).
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498
te
511
Trace essential elements
513
Within plant tissues, Fe has low solubility and high reactivity thus its movement along
514
the plant tissues must be associated with appropriate chelating compounds and proper
515
control of redox states (Kobayashi and Nishizawa, 2012). Several lines of evidence have
516
indicated that the main Fe chelators are citrate (Rellan-Alvarez et al., 2010, Harris et al.,
517
2012), NA (Ishimaru et al., 2010) and MAs (Aoyama et al., 2009, Kakei et al., 2012).
518
Citrate has been proved to play a key role in the Fe chelation and mobilization in xylem
519
sap. To facilitate the efflux of citrate into the xylem, Arabidopsis relies on the FRD3
520
(Ferric Reductase Defective 3) transporter which is expressed in the root pericycle cells
521
(Figure 5) (Durrett et al., 2007). In rice, an AtFDR3 orthologue, OsFRDL1, is also
522
expressed in root pericycle cells and encodes a citrate effluxer required to load citrate
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into the xylem sap (Yokosho et al., 2009). OsPEZ1 and OsPEZ2 are also responsible for
524
xylem loading of phenolics enhancing the remobilization of Fe along the plant body
525
(Bashir et al., 2011a, Ishimaru et al., 2011). Although AtFRD3, OsFRDL1, OsPEZ1 and
526
OsPEZ2 can efflux the Fe-chelating molecules, Fe must also be transported into the
527
xylem. To date, the plasma membrane-localized IREG1/FPN1 present in Arabidopsis
528
stele, is the only Fe effluxer identified to mediate the vascular loading of Fe (Morrissey
529
et al., 2009). Among the influx transporters, members of the YS and YSL transporters
530
are widely involved in Fe translocation. In barley and maize, the YS1 transporter is able
531
to mobilize Fe-PS complexes (Murata et al., 2008, Ueno et al., 2009). Among the 18
532
YSL transporters present in rice, OsYSL2, OsYSL15, OsYSL16, OsYSL18 were
533
identified as Fe-PS transporters involved in the Fe translocation. More specifically,
534
OsYSL2 is proved to be a Fe-NA transporter, whereas OsYSL15, OsYSL16, OsYSL18
535
translocate Fe-deoxymugineic acid (DMA) complexes (Aoyama et al., 2009, Inoue et
536
al., 2009, Ishimaru et al., 2010, Kakei et al., 2012). The efflux of NA in rice was found
537
to be facilitated by the ENA1 (efflux transporter of nicotianamine 1) and ENA2, and the
538
efflux of MA is achieved by TOM1 (Nozoye et al., 2011). Members of the YSL family
539
are also present in non-graminaceous plants that do not synthesize MAs. It is widely
540
assumed that non-graminaceous plants rely on the YSL transporters to translocate Fe-
541
NA complexes. NA is a precursor and structural analogue of MAs; it has the ability to
542
chelate Fe and other metals and is synthesized by all plants, in contrast to the
543
graminaceous-specific synthesis of MAs. The Arabidopsis YSL members AtYSL1,
544
AtYSL2 and AtYSL3 were identified to have a key role in the mobilization of Fe inside
545
the plant (Chu et al., 2010). Besides their presence in the outer root cells, some FRO
546
genes can also be expressed in root or shoot vascular cells (Jeong and Connolly, 2009),
547
suggesting that Fe reduction can occur within plant tissues with further Fe translocation
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by members of the IRT and/or NRAMP families (Takahashi et al., 2011). The
549
oligopeptide transporters (OPT) seem to play an important role in the mobilization of
550
Fe. OPT3 was markedly expressed in aerial parts of both Arabidopsis thaliana and
551
Thlaspi caerulescens, including stem and leaf, suggesting that this transporter may be
552
involved in the long-distance Fe transportation (Stacey et al., 2008, Hu et al., 2012).
553
In order to be translocated, Cu must be exported from the root symplast before entering
554
the xylem. Evidence suggests that the export from the root symplast is mediated by an
555
HMA family transporter. In Arabidopsis, AtHMA5 is highly expressed in roots, flowers
556
and pollen and seems to be responsible for root Cu detoxification (Andres-Colas,
557
Sancenon, 2006). This phenotype function is the opposite of COPT, corroborating the
558
idea that AtCOPT1 and AtHMA5 transport Cu in opposite directions (Figure 5) (del
559
Pozo, Cambiazo, 2010). Cu translocation may involve chelators such as NA and several
560
amino acids (Irtelli et al., 2009, Harris et al., 2012). Moreover, Cu in the xylem sap of
561
rice seems to be bound to DMA, while in the phloem sap, Cu mainly complexes with
562
NA and histidine (Ando et al., 2013). OsYSL16 is a Cu-NA transporter required for
563
delivering Cu to developing young tissues and seeds through phloem transport (Zheng
564
et al., 2012). In Arabidopsis root, the plasma membrane-localized AtYSL1 and AtYSL3
565
were proven to be involved in the root-to-shoot Cu transport (Waters and Grusak, 2008,
566
Chu et al., 2010). In Arabidopsis, AtCOPT6 is expressed in different cell types of
567
different plant compartments, but the bulk of its expression is located in the vasculature
568
suggesting a role in Cu translocation (Jung et al., 2012). Similarly, OsCOPT6 is not
569
expressed in root but is highly expressed in leaf, stem and sheath (Yuan et al., 2011).
570
Furthermore, the oligopeptide transporter AtOPT3 and its orthologue TcOPT3 seem to
571
participate in the mobilization of Cu (Stacey et al., 2008, Hu et al., 2012).
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The translocation of Mn involves chelators such as NA, amino acids and carboxylic
573
acids (Harris et al., 2012). In Arabidopsis, AtZIP2 seems to participate in Mn transport
574
into the root vasculature for translocation to the shoot (Figure 5) (Milner et al., 2013). In
575
rice, OsYSL6 is a Mn-NA transporter that is required for the detoxification of excess
576
Mn (Sasaki et al., 2011). Likewise, OsYSL2 is involved in Mn translocation in the
577
phloem. In the knockout ysl2 line, rice seeds and shoots contain lower Mn content
578
compared with the wild-type (Ishimaru et al., 2010). Moreover, the same transporter
579
(YSL2) shows the ability to transport Mn-PS complexes in barley roots (Araki et al.,
580
2011). The oligopeptide transporter AtOPT3 seems to play an important role in the
581
long-distance Mn transport (Stacey et al., 2008).
582
Nickel is translocated from roots to shoots as a free ion (Ni2+) and complexed with
583
several molecules such as histidine and citrate (Richau et al., 2009, Centofanti et al.,
584
2013). In hyperaccumulator plants, Ni was shown to be translocated also in the form of
585
Ni-NA complexes (Mari et al., 2006). The transport of NA-metal chelates, required for
586
the xylem loading and unloading, has been assigned to the YSL family. In Thlaspi
587
caerulescens, two YSL genes, TcYSL5 and TcYSL7 were found to be expressed around
588
the vasculature of the shoots and in the central cylinder of the roots, suggesting that
589
YSL transporters play a key role in the translocation of Ni-NA complexes (Gendre et
590
al., 2007).
591
Zinc is exported from root by two members of the P-type ATPase family, HMA2
592
(expressed in Arabidopsis, rice and barley) and HMA4 (Figure 5) (Mills et al., 2012,
593
Takahashi et al., 2012b). However, other transporters can also perform the export of Zn
594
to vascular tissues. In Arabidopsis, pcr2 knockout mutants accumulate Zn in roots,
595
suggesting a role of AtPCR2 in the root-to-shoot Zn translocation (Song et al., 2010).
596
Furthermore, it was proved that AtFRD3 is involved in loading Zn into xylem (Pineau
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et al., 2012). Zn can also be exported from the roots in the form of Zn complexes.
598
Before being exported, Zn must be mobilized to pericycle cells, apparently by HvYSL2
599
(Araki et al., 2011). After xylem loading, Zn can be transported to above-ground tissues
600
by the YS and YSL transporters. There is some evidence that AtYSL1, AtYSL2 and
601
AtYSL3 are involved in the transport of Zn (Chu et al., 2010). Phytosiderophores such
602
as NA and DMA are important in the distribution of Zn within the plant (Suzuki et al.,
603
2008, Harris et al., 2012, Nishiyama et al., 2012). Although no Zn-PS transporters have
604
been identified yet, the presence of these complexes in the xylem and phloem sap
605
suggests that members of the YSL family may be involved in the transport of Zn
606
complexes (Rellan-Alvarez et al., 2008, Deinlein et al., 2012). Moreover, members of
607
the ZIP family are expressed in the vascular bundle, suggesting that they contribute to
608
the distribution of Zn along the plant (Bashir et al., 2012). AtOPT3 and its orthologue
609
TcOPT3 seem to participate in the long-distance transport of Zn (Stacey et al., 2008, Hu
610
et al., 2012).
te
611
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597
Beneficial elements
613
The export of Na to the xylem is assumed to be an active process. In Arabidopsis, SOS1
614
was proposed to be involved in the loading of Na into the xylem (Shi et al., 2002).
615
However, at the whole-plant level, the specific role of AtSOS1 still remains uncertain
616
(Kronzucker and Britto, 2011). At high Na levels, xylem loading with Na is thought to
617
occur passively through channels, since a higher cytosolic Na content in xylem
618
parenchyma cells and a relatively depolarized plasma membrane would promote the
619
movement of Na into the xylem sap (Apse and Blumwald, 2007). Several lines of
620
evidence exist supporting the role of AtHKT1;1 and its orthologue OsHKT1;5 in xylem
621
unloading (Sunarpi et al., 2005, Davenport et al., 2007). A demonstration of Na efflux
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from shoot xylem vessels was achieved by overexpressing AtHKT1;1 in the mature root
623
stele of A. thaliana, which showed decreased Na accumulation in the shoot (Moller et
624
al., 2009).
625
For Co, little evidence is available regarding its translocation. However, an Arabidopsis
626
fpn1 knockout mutant showed low Co accumulation in shoot tissues, indicating that
627
ferroportins play an important role in Co homeostasis, more specifically in the root-to-
628
shoot Co translocation (Morrissey et al., 2009).
us
629
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ip t
622
Leaves compartmentation
631
The transport of inorganics into leaf cells from leaf xylem involves another membrane
632
transport step mediated by specific membrane transport proteins. Once inside the leaf
633
symplast, cations can be assimilated in the cytosol or stored in cellular organelles. The
634
general rule is that nutrients are assimilated, playing a key role in several physiological
635
mechanisms, and toxic elements are stored in places where they cause the least harm to
636
cellular processes. However, it should be noted that nutrients can also be stored for two
637
reasons: (i) they can become deficient and plant accumulates them for times of
638
necessity; (ii) they are at high concentrations and can pose a toxicity problem to plants
639
(Maathuis, 2009, Pilon-Smits et al., 2009, Pilon et al., 2009). At the tissue level, toxic
640
elements are generally accumulated in the epidermis and trichomes (Tian et al., 2009,
641
Sanchez-Pardo et al., 2012); at the cellular level, they may be accumulated in the
642
vacuoles or cell wall (Cosio et al., 2005).
643
The distribution of cations in leaf tissues is generally asymmetric (Wu and Becker,
644
2012). The accumulation of toxic cations in leaf vacuoles makes sense because vacuoles
645
do not contain a photosynthetic apparatus, which would be sensitive to toxicity
646
(Martinoia et al., 2012). Toxic cation storage occurs also in highly tolerant cells such as
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the leaf epidermis and the vein bundle sheath, as long as their import remains under
648
control. Once the cation accumulation exceeds the tolerance threshold of the plant, they
649
would be transported to mesophyll cells, which are more sensitive to toxic cations than
650
other cell types, allowing photosynthesis to be threatened (Zhao et al., 2012). At
651
present, the physiological mechanisms involved in the sequestration of cations between
652
different leaf tissues remain only partially understood. One possible mechanism is the
653
differential expression of cation transporters in the plasma membrane and/or tonoplast
654
between mesophyll and other tissues (Morel et al., 2009).
us
cr
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647
an
655
Macroelements
657
The transport of K from the xylem into the leaf symplast is mediated by membrane
658
transporters and channels. Members of the HAK/KT/KUP family were found to be
659
highly expressed in shoot tissues. Specifically, AtKUP1, AtKUP2 and AtKUP4 were
660
expressed in Arabidopsis leaves, suggesting an involvement in K xylem unloading
661
(Wang and Wu, 2013). Moreover, Ahn et al (2004) also showed that other AtKUPs
662
were highly expressed in leaves. In particular, AtKUP6, AtKUP8 and AtKUP9 were
663
highly expressed in young leaves, whereas AtKUP10 and AtKUP12 expression was
664
evident in both older and younger leaves. Shaker channels AtKT1, AtKT2 and AtKC1
665
are also highly expressed in the plasma membrane of Arabidopsis leaves, indicating that
666
these cation channels are enrolled in the movement of K between shoot tissues
667
(Jeanguenin et al., 2011). Regarding the intracellular distribution of K in leaves, the
668
vacuolar K transporters AtNHX1 and AtNHX2, expressed in the tonoplast of
669
Arabidopsis leaf vacuoles, mediate the influx of K into this cell organelle (Figure 6)
670
(Bassil et al., 2011). To mediate the vacuolar K efflux, several members of the
671
TPK/KCOs family, such as AtTPK1, AtTPK3, AtTPK5 and AtKCO3, are expressed in
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the vacuolar membrane of leaf vacuoles and mediate this process (Voelker et al., 2010,
673
Hamamoto and Uozumi, 2014). Regarding other cell organelles, a member of the CHX
674
family, AtCHX23, was found to be expressed in the chloroplast envelope of
675
Arabidopsis leaf cells. Analysis of Arabidopsis chx23 knockout mutant leaves suggests
676
that CHX23 could mediate the co-transport of K across the chloroplasts envelope of leaf
677
cells (Song et al., 2004). Recently, a member of the K efflux antiporters (KEA) has also
678
been identified in Arabidopsis chloroplasts. AtKEA2 was found to be highly expressed
679
in leaves and mediates K transport across the plastid envelope (Aranda-Sicilia et al.,
680
2012).
681
Within the leaf, Ca follows mainly the apoplastic pathway and tends to accumulate in
682
the mesophyll cells, trichomes and epidermal cells. The cytosolic Ca concentration is
683
very sensitive to the apoplastic Ca concentrations. Thus, Ca is usually stored in leaf
684
vacuoles and/or specific cell organelles to avoid excessive apoplastic Ca accumulation
685
(White and Broadley, 2003). However, before been sequestered in the leaf vacuoles, Ca
686
must be transported across the plasma membrane of leaf cells. Although most members
687
of the NSSC are present in the root cells, several lines of evidence suggest that VI-
688
NSCCs can also be found in leaf cells (Demidchik and Maathuis, 2007). In the vacuoles
689
of leaf cells, Ca is present as the ionic species Ca2+, in soluble forms (e.g., complexed
690
with proteins and/or organic acids) and/or in insoluble forms (e.g., Ca-oxalate and Ca-
691
phytate) (Karley and White, 2009). The transport of Ca across the vacuolar membrane
692
of leaf cells is thought to be performed by members of the ACA and CAX families
693
(Figure 6). AtACA11 is targeted to the vacuolar membrane of Arabidopsis leaf cells and
694
mediates Ca transport across the tonoplast of large vacuoles (Lee et al., 2007). In the
695
CAX family, AtCAX1 is highly expressed in the vacuoles of leaf cells and has an
696
important role in Ca influx into the vacuole (Cheng et al., 2005, Conn et al., 2011b).
Ac ce p
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29 Page 29 of 78
AtCAX2, AtCAX5 and AtCAX6 were also expressed in Arabidopsis leaves, although in
698
a lower extent than AtCAX1 (Edmond et al., 2009).
699
Members of the MGT/MRS2 family are thought to mediate the leaf uptake of Mg
700
(Gebert et al., 2009, Karley and White, 2009). It was shown that AtMGT9 is expressed
701
in leaf cells and thus can be involved in Mg transport (Chen et al., 2009). Once in the
702
cytosol of leaf cells, Mg can be accumulated in several cell organelles. Evidence
703
suggests that mesophyll cells accumulate the highest vacuolar concentration of Mg in
704
Arabidopsis leaves. To accomplish that, plants rely on members of the MGT/MRS2
705
family, such as AtMGT2/MRS2-1 and AtMGT3/MRS2-5, which are localized in the
706
tonoplast of Arabidopsis leaf cells and mediate the Mg influx into the vacuole (Figure
707
6) (Conn et al., 2011a). AtMHX is also involved in Mg influx across the tonoplast of
708
leaf cells (Shaul et al., 1999, Elbaz et al., 2006). A member of the MGT/MRS2 family,
709
AtMRS2-11, was expressed in the chloroplast envelope and plays a key role in the Mg
710
transport into this cell organelle (Drummond et al., 2006).
cr
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711
ip t
697
Trace essential elements
713
Fe can be transported into the leaf symplast as free ion or complexed with PS. Members
714
of the IRT family are expressed in leaf tissues of Arabidopsis suggesting their
715
involvement in the Fe acquisition (Lee and An, 2009, Lin et al., 2009, Barberon et al.,
716
2011). Regarding Fe-PS complexes, members of the YSL family seems to mediate the
717
Fe influx into the cytosol of leaf cells (Kakei et al., 2012). Once in the leaf symplast, Fe
718
must be delivered into leaf cells to participate in multiple biochemical processes. The
719
chloroplast is the largest pool of Fe in plant cells accumulating 80 to 90% of cellular Fe
720
(Kobayashi and Nishizawa, 2012). The influx of Fe into the chloroplast of Arabidopsis
721
thaliana is achieved by the PIC1 (permease in chloroplast 1) localized in the inner
Ac ce p
712
30 Page 30 of 78
envelope membrane of chloroplasts (Figure 7) (Duy et al., 2011). AtFRO7 is localized
723
in the chloroplast envelope and looks to be critical for chloroplast Fe acquisition (Jeong
724
and Connolly, 2009). Mitochondria are other plant sites where Fe is essential. Bashir et
725
al. (2011b) identified the rice mitochondrial Fe transporter (MIT), which is essential for
726
rice growth and development. Moreover, AtFRO3 and AtFRO8 are localized in
727
mitochondria membrane, which indicates that these ferric reductase oxidases must be
728
involved in mitochondrial Fe homeostasis (Jeong and Connolly, 2009). The vacuole
729
generally functions as a metal pool to avoid toxicity. The transporters OsVIT1 and
730
OsVIT2 mediate the influx of Fe across the vacuolar membrane into the vacuole (Zhang
731
et al., 2012). Likewise, it seems that the IREG2/FPN2 present in Arabidopsis tonoplast
732
may play a role in the influx of Fe from the cytoplasm into the vacuole (Morrissey et al.,
733
2009). On the other hand, both AtNRAMP3 and AtNRAMP4 export Fe from vacuoles
734
(Lanquar et al., 2005). In the Golgi complex, the MATE (multidrug and toxic
735
compound extrusion) transporter BCD1 has a role in sustaining Fe homoeostasis by
736
reallocating excess Fe released from stress-induced cellular damage in Arabidopsis (Seo
737
et al., 2012).
738
In order to Cu reach the leaf symplast, plasma membrane proteins must exist to mediate
739
Cu xylem unloading and Cu acquisition. It seems that members of the COPT family are
740
responsible for the influx of monovalent Cu (Sancenon et al., 2004, Andres-Colas et al.,
741
2006). However, since Cu is present mainly in the divalent form in the vascular tissues,
742
reduction must occur before it can enter the cell organelles. AtFRO7 (expressed in the
743
chloroplasts) and AtFRO3/8 (expressed in the mitochondria) play a central role in Fe
744
reduction, and it is hypothesized that they also participate in Cu reduction (Jeong and
745
Connolly, 2009). Moreover, Cu can also be transported across the plasma membrane of
746
leaf cells complexed with PS. Members of the YSL family were proved to be engaged
Ac ce p
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722
31 Page 31 of 78
in the Cu translocation and delivery to shoot tissues (Irtelli et al., 2009, Zheng et al.,
748
2012, Ando et al., 2013). Once in the cytosol of leaf cells, Cu is distributed with the aid
749
of plasma membrane-localized transporters among different cell organelles. AtHMA6
750
(or PAA1), localized in the inner chloroplast envelope, is responsible for the delivery of
751
Cu to chloroplasts (Figure 7) (Catty et al., 2011). AtHMA8 (PAA2), closely related to
752
AtHMA6 (PAA1), is expressed in the thylakoid membrane and supplies Cu to
753
plastocyanin (Tapken et al., 2012). AtHMA1 and HvHMA1, present in the chloroplast
754
envelope, are broad-specificity exporters of metals from chloroplasts and may play a
755
specialized role in Cu mobilization (Mikkelsen et al., 2012). AtHMA7 (RAN1), the first
756
functionally characterized heavy metal ATPase, is responsible for the biogenesis of
757
ethylene receptors by supplying Cu at the endoplasmic reticulum and also for Cu
758
homeostasis in seedling development (Binder et al., 2010). The rice mitochondrial Fe
759
transporter (MIT) also appears to regulate the influx of Cu, although more studies are
760
needed to confirm this (Bashir et al., 2011b). The tonoplast-localized AtCOPT5 is
761
important for Cu export from the vacuole and is involved in the remobilization of Cu
762
ions (Garcia-Molina et al., 2011, Klaumann et al., 2011).
763
Manganese reaches the leaf symplast by crossing the plasma membrane of leaf cells. To
764
do that, members of the IRT family are thought to be expressed in the shoot tissues,
765
mediating the transport of Mn ions into the cytosol of leaf cells (Pedas et al., 2008,
766
Barberon et al., 2011). After influx into the leaf symplast, Mn must be distributed into
767
various cell compartments. In Arabidopsis, AtECA1 is localized at the endoplasmic
768
reticulum (Li et al., 2008). The cation/H+ exchanger (CAX) transporters AtCAX2 and
769
AtCAX4, which were originally identified as Ca2+ transporters, also have the ability to
770
transport Mn into the vacuole (Figure 7) (Connorton et al., 2012). Likewise, vacuolar
771
Mn2+/H+ antiporter activity in the Arabidopsis cax2 knockout mutant is significantly
Ac ce p
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747
32 Page 32 of 78
reduced, although it is not completely absent, compared with wild-type, suggesting the
773
presence of additional vacuolar transporters (e.g., AtCAX5) that contribute to Mn
774
transport (Edmond et al., 2009). AtMTP11, which belongs to the cation diffusion
775
facilitator (CDF) family, is implicated in the pre-vacuolar compartmentation of Mn as
776
well as in Mn homeostasis mechanisms (Delhaize et al., 2007). Other MTPs also
777
contribute to Mn transport (Gustin et al., 2011). Furthermore, it seems that the
778
tonoplast-localized transporters OsVIT1 and OsVIT2 participate in Mn influx to the
779
vacuole (Zhang et al., 2012). Regarding vacuolar export, AtNRAMP3 and AtNRAMP4
780
are expressed in the tonoplast and can transport Mn and other metals (Lanquar et al.,
781
2010). AtZIP1 probably plays a role in Mn vacuolar efflux, based on the increased
782
sensitivity to low Mn and increased accumulation of Mn in roots in the zip1 knockout
783
line (Milner et al., 2013). The rice Fe transporter MIT appears to play a role in Mn
784
transport into mitochondria (Bashir et al., 2011b). The mechanisms of Mn transport in
785
the chloroplast remain unknown, despite the fact that Mn has a critical role in
786
photosynthesis and the chloroplast is the one of the major sinks for Mn (Yao et al.,
787
2012, Millaleo et al., 2013).
788
Zinc enters the leaf symplast with the aid of members of the IRT and ZIP families
789
(Sinclair and Kramer, 2012). After reaching the leaf cytosol, the delivery of Zn to plant
790
organelles is also mediated by specific transporters (Figure 7). The vacuolar membrane
791
transporters AtMTP1 and AtMTP3 are expressed in the vacuoles of leaf cells and play a
792
key role in detoxification of excessive Zn by accumulating Zn in vacuoles (Arrivault et
793
al., 2006, Gustin et al., 2009). Likewise, AtHMA3 is also involved in vacuolar Zn
794
sequestration (Morel et al., 2009). In Arabidopsis, AtZIF1 also seems to be expressed in
795
the tonoplast of leaf cells. Therefore, an active role is expected from this transporter
796
regarding the vacuolar Zn sequestration (Haydon et al., 2012). The tonoplast-localized
Ac ce p
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772
33 Page 33 of 78
OsVIT1 and OsVIT2 seem to transport Zn into the vacuoles of plant cells (Zhang et al.,
798
2012). In the chloroplasts of both Arabidopsis and barley, HMA1 contributes to Zn
799
detoxification by exporting Zn from the plastids to the cytoplasm (Kim et al., 2009,
800
Mikkelsen et al., 2012). It has been proposed that the Golgi-localized transporter
801
AtECA3 may have a role in the mobilization of Zn from the cytoplasm to the Golgi
802
complex (Mills et al., 2008).
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803
Beneficial elements
805
Regarding the beneficial cations Na and Co, little information is available about the
806
processes regulating their movement into the leaf symplast as well as their distribution
807
in the cell organelles (Pilon-Smits et al., 2009). Although their essentiality is not yet
808
proved, evidence suggests that similar membrane transporters should mediate their
809
mobilization within the plant compartments (Hänsch and Mendel, 2009, Kronzucker
810
and Britto, 2011).
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811
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804
Biofortification
813
Plant foods refer to plants or parts of a plant that are eaten as a food. In this category are
814
included vegetables, fruits, grains and legumes. They constitute one of the most
815
important nutrient sources in human diet since the beginning of civilization. Humans
816
need several essential minerals in order to allow the proper functioning of the body,
817
regarding growth, development and metabolism. Recommended dietary allowances
818
(RDAs), defined as “the levels of intake of essential nutrients that, on the basis of
819
scientific knowledge, are judged by the Food and Nutrition Board to be adequate to
820
meet the known nutrient needs of practically all healthy persons” are established and
821
vary between a few μg/day and 1 g/day. If intakes are low in a certain period of time,
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deficiency signs may develop. On the other hand, high intakes can result in toxicity
823
(FNB/IOM, 2006).
824
To grow and develop, plants must take up water and essential mineral nutrients from the
825
soil. Once taken up by plant roots, nutrients can be transported and accumulate in other
826
parts of the plant. The final composition of the edible parts of plants is influenced and
827
controlled by several factors, including soil fertility, plant genetics and the environment
828
in which they grow (Karley and White, 2009, Maathuis, 2009, Puig and Penarrubia,
829
2009). Although it is possible in some cases to enhance the nutritional quality of plant
830
foods through agronomic practices and plant breeding, the soil-to-plant and plant-to-
831
human transfer of mineral nutrients is a very complex process (Hirschi, 2009). First of
832
all, soils differ greatly in their mineral composition and nutrients phytoavailability
833
depends on several factors (Palm et al., 2007, Pinto et al., 2014b). Secondly, plants
834
possess physiological mechanisms to regulate the amounts of nutrients taken up from
835
the soil. Hence, one can expect that attempts to alter the overall mineral composition of
836
plant foods will be very complex (Maathuis, 2009, Pilon et al., 2009, Pinto et al.,
837
2014a). Thirdly, it has been recognized that the concentration of a nutrient in a food is
838
not necessarily a reliable indicator of the value of that food as a source of the nutrient in
839
question. This leads us to the concept of nutrient bioavailability, defined as the amount
840
of a nutrient in ingested food that is available for utilization in metabolic processes. In
841
the case of mineral nutrients, bioavailability is determined mainly by the efficiency of
842
minerals absorption from the human gut (Hirschi, 2009). Therefore, the development of
843
plant foods biofortification must rely on a holistic approach focusing on three concepts:
844
(1) nutrients phytoavailability, (2) nutrients accumulation in edible tissues and (3)
845
nutrients bioavailability. In this review, only the two first steps (nutrients
846
phytoavailability and nutrients accumulation in edible tissues) will be discussed. For
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35 Page 35 of 78
847
information regarding nutrients bioavailability, the reader should refer to the work of
848
Hirschi (2009).
849
Nutrients phytoavailability
851
Five areas have been identified that are related to the uptake efficiency of cations by
852
plants in which breeders can act. These include: (1) root morphology, (2) root hairs
853
formation, (3) root exudates, (4) ability to release cations from non-exchangeable pools,
854
and (5) kinetics of cation uptake. Thus, a successful biofortification strategy must
855
understand three important aspects: (1) the spatial distribution of elements within soil
856
profiles, (2) in which form elements are commonly found in the soil, and (3) how plant
857
roots acquire these elements. Gathering this knowledge is of utmost importance and will
858
enable researchers to concentrate efforts in the most limiting steps of nutrient
859
acquisition.
860
Most cations in soil are incorporated in the crystal lattice structure of minerals and thus
861
not readily available for plant uptake. The availability of these elements will be greatly
862
influenced by the soil type and its physico-chemical properties. To simplify, cations in
863
the soil can be classified in four groups depending on their phytoavailability: (1) water-
864
soluble; (2) exchangeable; (3) non-exchangeable and (4) structurally bound. Water-
865
soluble cations are directly available for both plants and microorganisms and are highly
866
mobile within soil profiles. Exchangeable cations are those weakly adsorbed elements
867
retained on the solid surface by relatively weak electrostatic interaction as well as
868
cations that can be released by ion-exchange processes (Fedotov et al., 2012). Both
869
water-soluble and exchangeable pools are usually considered to be easily
870
phytoavailable. However, the size of both pools is negligible, accounting for as much as
871
0.1-0.2% and 1–2% of the total content of these elements in soil, respectively. Non-
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36 Page 36 of 78
exchangeable and structurally bound pools are considered to be slowly or even non-
873
available sources of elements for plants. These pools can only contribute to the supply
874
of nutrients to plants in the long term. (Zörb et al., 2014). As already described, the
875
amount of cations that are phytoavailable and non-phytoavailable in the soil vary greatly
876
among soil types. However, in all soils a dynamic equilibrium exist between the
877
different pools, which is influenced by a number of soil physico-chemical properties as
878
well as plant-soil interactions.
879
The main soil physico-chemical properties that control the processes of sorption and
880
desorption of elements in the soil are: (1) pH, (2) cation exchange capacity (CEC), (3)
881
particle size distribution, mainly the fine granulometric fraction (< 0.02 mm), (4)
882
organic matter (OM) and (5) oxides and hydroxides, mainly of Al, Fe and Mn (Kabata-
883
Pendias, 2004). Typically, the most mobile pool of elements occur at lower pH. CEC
884
reflects the dynamic equilibrium of cationic species in the soil solution, thus influencing
885
the phytoavailability of elements. When soil pH becomes acid, the phytoavailability of
886
elements usually increases due to the replacement of cations on soil binding sites by H+
887
ions. Clay minerals and oxides usually possess high active surface areas, which confer
888
them the ability to strongly bind cations. OM increases the CEC of soils because OM
889
possesses multiple binding sites for cations, which results in a great sorption capacity
890
(Pinto et al., 2014a).
891
With respect to plant-soil interactions, the rhizosphere processes such as root exudation,
892
nutrient mobilization and rhizosphere respiration, are known to greatly influence the
893
phytoavailability of elements (Hinsinger et al., 2009). Root exudation includes the
894
secretion of ions, free oxygen, water, enzymes, mucilage, and a wide variety of primary
895
and secondary metabolites (Badri and Vivanco, 2009). These exudates can increase or
896
decrease the availability of nutrients by altering soil physico-chemical properties and
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37 Page 37 of 78
soil biological processes. Typically, these root-induced changes will alter the solubility
898
and speciation of elements in the rhizosphere in a way that nutrients are more easily
899
available for plant uptake (Badri and Vivanco, 2009, Kobayashi and Nishizawa, 2012).
900
It is now clear that one of the ways to develop more efficient plant foods, especially in
901
terms of the nutrient density, is by improving the root exudation process. This could
902
shift the equilibrium towards the enhance contribution of non-exchangeable and
903
structurally bound pools to the supply of nutrients that are many times not available for
904
plant uptake and also promote root growth as well as the formation of root hairs.
905
Breeders can start looking at the genes encoding the plasma membrane H+-ATPase
906
(HA), which are responsible for the efflux of protons that promote the acidification of
907
the rhizosphere. For instance, the breeding for an increased H+ efflux from the root in
908
order to promote the acidification of the surrounding soil may contribute to increase the
909
availability of virtually any element required for plant growth since, as above
910
mentioned, almost all elements are more mobile and thus phytoavailable at low soil pH.
911
Furthermore, H+ excretion is essential in maintaining or promoting primary root
912
elongation and root hair development (Palmgren, 2001a). Some attempts have been
913
already made in this area. In Arabidopsis, the protein kinase 5 (PKS5) and the
914
chaperone DNAJ homolog 3 (J3) have been identified to play important roles in the
915
efflux of H+ by regulating the interaction between the plasma membrane HA and 14-3-3
916
proteins (Yang et al., 2010). Furthermore, PIN2 (an auxin efflux transporter) was also
917
identified to be involved in the modulation of H+ secretion in the root tip, maintaining
918
primary root elongation (Xu et al., 2012a). More recently, it has been demonstrated the
919
involvement of 14-3-3 proteins in the regulation of H+ efflux and primary root
920
elongation. Gene specificity and functional redundancy was observed among 14-3-3
921
proteins in primary root elongation under both control and abiotic stress conditions (van
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38 Page 38 of 78
Kleeff et al., 2014). Furthermore, TFT4 and TFT7 (tomato 14-3-3 proteins), separately
923
introduced into Arabidopsis and overexpressed, showed enhanced plasma membrane
924
HA activity, H+ efflux and also maintains elongation of the primary root (Xu et al.,
925
2012b, Xu et al., 2013a). It has been also shown that abcissic acid accumulation
926
modulates auxin transport in the root tip, which further enhances H+ secretion (Xu et al.,
927
2013b).
928
Another way to improve the nutrient density of plant foods is to manipulate the root
929
system architecture towards a distribution of roots in the soil that optimizes nutrients
930
uptake. It is important that plants have a large surface area of contact between roots and
931
soil. This allows them to have more “options” to get the water and nutrients they need.
932
The growth of the main and lateral roots is supported by cells produced in the root
933
apical meristem. In this process, phytohormones play a key role. Auxin and cytokinin
934
are assumed to be the two most important regulators of root development (Jones and
935
Ljung, 2012). The presence of auxin concentration gradients with a maximum centered
936
at the quiescent center were revealed to be essential for the establishment and
937
maintenance of the stem cell niche and for transit amplifying cell divisions (Petersson et
938
al., 2009). Cytokinin acts as an antagonist of auxin. In fact, the balance between root
939
division and differentiation appears to be primarily maintained by this antagonistic
940
relationship, with cytokinin limiting the size of the root apical meristem and the rate of
941
root growth (Zhang et al., 2011). Lateral roots are formed from pericycle cells
942
adjacently located to the xylem poles. Several genes seem to be involved in this process
943
(Moreno-Risueno et al., 2010). The identification of candidate genes will ultimately
944
allow breeders to manipulate the root system architecture to facilitate the search for
945
water and nutrients by plants. Efficient genotypes could be develop to have a relatively
946
larger proportion of thin roots as well as high number of root hairs in the whole root
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39 Page 39 of 78
system. This is thought to enhance the phytoavailability and uptake of nutrients required
948
for plant growth and development.
949
For graminaceous plants, the overexpression of genes responsible for the secretion of
950
PS will have a great impact in the solubilization of trace elements such as Fe and Zn
951
that are usually low on plant foods (Nozoye et al., 2011). Candidate genes encoding
952
ENA1 and ENA2 could be manipulated to enhance the efflux of NA in graminaceous
953
plants such as rice. Similarly, plants overexpressing TOM1 will increase the efflux of
954
MA, which could result in the increase solubilization of trace elements for further
955
uptake by plant roots.
956
Despite these improvements, it may be also necessary to manipulate the plant uptake
957
system. If increased uptake of nutrients by roots is an important goal for breeding of
958
efficient plant foods, researchers may need to take a look on the role of each nutrient in
959
plant’s metabolism. For instance, the dominant role of K is the osmoregulation. Its role
960
in turgor provision and water homeostasis is obvious in processes such as pressure-
961
driven solute transport in the xylem and phloem and high levels of vacuolar K
962
accumulation (Wang and Wu, 2013). Thus, the breeding for an increased activity of K
963
transporters/channels (e.g., AtKT/KUP or AKT1) may contribute to an increase in cell
964
turgor, leading to cell elongation during the growth of root hairs, conferring advantages
965
to plant foods growth under limiting conditions (Nieves-Cordones et al., 2014).
966
However, care should be taken to properly assess the influence of overexpressing lines
967
in maintaining the charge balance within the plant cells, which is crucial for plant
968
development. To cope with an increase in cation uptake, plants have also to increase the
969
uptake of anions like chloride, nitrate, phosphate or sulphate that also act as osmolytes.
970
This will maintain the homeostasis within plant cells. Overall, when breeding is a goal
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40 Page 40 of 78
971
for enhance nutrient density of plant foods, ionomic studies at the whole plant level
972
should be carried out to assess the elemental distribution within plant cells.
973
Nutrients accumulation in edible tissues
975
The successful allocation of nutrients in the edible tissues of a certain plant greatly
976
depends on the plant tissue that is considered “edible”. For instance, the edible part of
977
carrots and potatoes are the roots. Therefore, in this case, attempts should be made to
978
enhance the accumulation of essential nutrients in the root of these plants. By contrast,
979
the edible tissues of leafy vegetables (e.g., lettuce or spinach) are the stems and leaves.
980
Thus, nutrients translocation must occur to move nutrients to above-ground tissues.
981
Finally, in other cases, seeds are the edible tissues of certain plants (e.g., rice) and
982
nutrients loading into seeds need to occur.
983
Two areas have been identified that are related to the utilization efficiency of elements
984
by plants in which breeders can act. These include: (1) cation translocation, and (2)
985
cation sequestration/storage. Some attempts have already been made in the two above-
986
mentioned areas. To improve cation translocation, a P1B-ATPase (AtHMA4) involved in
987
the transport of Zn in Arabidopsis was expressed in tobacco, showing enhanced Zn
988
translocation to the shoot (Siemianowski et al., 2011). Furthermore, the ectopic
989
expression of the same export protein (AtHMA4) in tobacco also resulted in decreased
990
Cd uptake/accumulation in roots and shoots (Siemianowski et al., 2014). In addition to
991
the main aim of enhancing the nutrient density of plant foods, these findings will also
992
allow breeders to manipulate plants in order to reduce the toxic exposure of humans via
993
intake of plant foods-derived Cd. Similar results have been obtained for other export
994
proteins involved in the translocation of other cations to above-ground tissues
995
(Maathuis, 2009, Pilon-Smits et al., 2009, Pilon et al., 2009, Kobayashi and Nishizawa,
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41 Page 41 of 78
2012, Mills et al., 2012, , Takahashi et al., 2012a, Ahmad and Maathuis, 2014). To
997
improve cation sequestration/storage in the organelles of edible plant tissues, breeders
998
may try to manipulate vacuolar transporters. It is now widely accepted that increasing
999
the capacity of edible tissues to sequester nutrients can promote their accumulation in
1000
these tissues. Thus, promoting the activity of vacuolar influx transporters, such as VIT,
1001
HMA, MTP and/or IREG2/FPN2, in above-ground edible tissues is seen as a promising
1002
way of food biofortification. Some research have already been made in this field. For
1003
instance, transgenic carrot and potato overexpressing the Arabidopsis vacuolar H+/Ca2+
1004
transporter AtCAX1 showed increased Ca content compared to the wild-type (Park et
1005
al., 2004, Park et al., 2005). In a similar way, potato tubers expressing the A. thaliana
1006
vacuolar transporter AtCAX2B [a variant of N-terminus truncated form of CAX2
1007
(sCAX2)] accumulate two times more Ca than wild-type (Kim et al., 2006a). Similar
1008
attempts have also been made to enhance the nutrients density of leaves and seeds. For
1009
instance, it has been shown that overexpression FRO genes results in an increase
1010
accumulation of Fe in plant leaves (Jeong and Connolly, 2009). Additionally, two
1011
important rice genes, OsVIT1 and OsVIT2, have been recently identified and
1012
characterized. It has been shown that functional disruption of both genes leads to
1013
increased Fe/Zn accumulation in rice seeds, which might represent another strategy to
1014
biofortify Fe/Zn in staple foods (Zhang et al., 2012). However, with respect to Fe, it is
1015
also necessary to increase the biosynthesis of Fe-chelates in the shoot to increase Fe
1016
concentrations in seeds and/or fruits, which rely on Fe supplied via the phloem
1017
(Aoyama et al., 2009, Inoue et al., 2009, Morrissey et al., 2009, Ishimaru et al., 2010,
1018
Kakei et al., 2012).
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1019 1020
Final Remarks 42 Page 42 of 78
The main roles of essential elements have been well documented and they are unlikely
1022
to change. However, many aspects related to whole-plant nutrition needs further
1023
elucidation. For instance, some studies have already described a shift in mineral
1024
nutrition in overexpressing or knockout lines of a certain membrane transporter.
1025
Mutations that increased the content of certain elements in plant tissues usually showed
1026
negative pleiotropic effects in the content of other essential mineral elements and even
1027
on plant growth. It is now widely recognized that a complex network exists between
1028
elements and that biotic and abiotic stress are likely to change not only one element but
1029
the all ionomic profile of a plant. Despite the great advance achieved in last years,
1030
further studies are required to clarify some aspects of plant nutrition. The search for new
1031
players in the field of cation transport still represents a challenge, particularly for trace
1032
essential and beneficial elements. Evidence suggests that the beneficial elements Al, Co
1033
and Na have positive effects on plant growth and stress resistance. Despite that, the
1034
mechanisms behind the beneficial effects of these elements to plants are relatively
1035
underinvestigated, especially when compared to the toxic effects that many of these
1036
elements have at high levels. It is clear that more attention should be given to the effects
1037
of beneficial elements at low levels, since this will not only clarify several basic
1038
processes of plant nutrition but also because fertilizing with beneficial nutrients may
1039
improve crop production and affect plant nutritional value.
1040
This review summarizes the most recent discoveries regarding the function and
1041
regulation of the multiple systems involved in cation transport in plant cells. Since most
1042
of the work has been performed with the model plant Arabidopsis thaliana, the next
1043
step should be the transfer of such knowledge to crop species. The specific properties,
1044
regulation and/or expression patterns of cation transporters/channels in different species
1045
may account for relevant agronomic advances. It is expected that in the upcoming years
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many promising results will be achieved in the field of biofortification. However, the
1047
application of biofortification programs will ultimately depend on the cost/benefit
1048
evaluation that farmers, consumers and politics will do. In addition, one of the most
1049
important tasks that researchers need to assure is the careful and systematic analysis of
1050
these foods before they become available to consumers. In this way, it seems that
1051
‘omics’ methods will have a crucial role providing valuable information regarding the
1052
actual safety of these foods.
cr
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1046
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1053
44 Page 44 of 78
1053
Figures Caption
1055
Fig. 1 – Schematic illustration of some of the key transport steps for the uptake of K, Ca
1056
and Mg in the plant (Arabidopsis thaliana was used as model plant). The proteins
1057
identified for mediating some of these transport functions are given in the table below.
1058
The arrows indicate the direction of transport and each letter in the figure corresponds to
1059
the list given in the first column of the table.
1060
Fig. 2 – Fe acquisition strategies in plants: (left) Strategy I in non-graminaceous plants
1061
and (right) Strategy II in graminaceous plants (right). Black boxes represent the
1062
transporters and enzymes that play essential roles in these strategies
1063
Fig. 3 – Schematic illustration of some of the key transport steps for the uptake and
1064
reduction of Cu Mn, Ni and Zn in the plant (Arabidopsis thaliana, Zea mays and
1065
Hordeum vulgare were used as model plants). The proteins identified for mediating
1066
some of these transport/reductive functions are given in the table below. The arrows
1067
indicate the direction of transport/reduction and each letter in the figure corresponds to
1068
the list given in the first column of the table.
1069
Fig. 4 – Tolerance mechanisms for cations in plant cells. Detoxification usually
1070
involves conjugation followed by active sequestration in the vacuole and apoplast,
1071
where the toxic cations can be less harmful. Chelators shown are GSH: glutathione,
1072
MT: metallothioneins, NA: nicotianamine, OA: organic acids, PC: phytochelatins.
1073
Active transporters are shown as black boxes with arrows.
1074
Fig. 5 – Schematic illustration of some of the key transport steps for the translocation of
1075
Fe, Cu Mn and Zn in the plant (Arabidopsis thaliana and Oryza sativa were used as
1076
model plants). The proteins identified for mediating some of these transport functions
Ac ce p
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45 Page 45 of 78
are given in the table below. The arrows indicate the direction of transport and each
1078
letter in the figure corresponds to the list given in the first column of the table.
1079
Fig. 6 – Schematic illustration of the subcellular localization of transporters involved in
1080
the movement of K, Ca and Mg in Arabidopsis thaliana. Members of the AtNHX,
1081
AtTPK, AtKCO, AtCHX, AtKEA, AtACA, AtCAX, AtMGT, AtMHX and AtMRS
1082
families are targeted in intracellular membranes: tonoplast (AtNHX1-2, AtACA11,
1083
AtCAX1, AtMGT2, AtMGT3, AtMHX, AtMRS2-11, AtTPK1, AtTPK3, AtTPK5 and
1084
AtKCO3) and chloroplast envelope (AtCHX23 and AtKEA2). The arrows indicate the
1085
direction of transport and each letter in the figure corresponds to the list given in the
1086
first column of the table.
1087
Fig. 7 – Schematic illustration of the subcellular localization of transporters and
1088
reductases involved in Fe, Cu, Mn and Zn movement and reduction, respectively, in
1089
Arabidopsis thaliana and Oryza sativa. Members of the OsVIT, AtFPN, AtNRAMP,
1090
AtBCD, AtPIC, AtFRO, OsMIT, AtCOPT, AtHMA, AtCAX, AtMTP, AtZIP, AtZIF
1091
and AtECA families are targeted in intracellular membranes: vacuole (OsVIT1/2,
1092
AtFPN2, AtCAX2/4-5, AtMTP1/3/11, AtZIF1, AtNRAMP3/4, AtCOPT5 and AtZIP1),
1093
Golgi complex (AtBCD1 and AtECA3), chloroplast (AtPIC1, AtHMA1/6 and
1094
AtFRO7), thylakoid membranes (AtHMA8), mitochondrion (OsMIT and AtFRO3/8)
1095
and endoplasmic reticulum (AtHMA7). The arrows indicate the direction of
1096
transport/reduction and each letter in the figure corresponds to the list given in the first
1097
column of the table.
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1098
46 Page 46 of 78
1098
1101 1102 1103
Ahmad I, Maathuis FJM. Cellular and tissue distribution of potassium: Physiological relevance, mechanisms and regulation. Journal of Plant Physiology. 2014;171:708-14. Ahn SJ, Shin R, Schachtman DP. Expression of KT/KUP genes in arabidopsis and the role
ip t
1100
REFERENCES
of root hairs in K+ uptake. Plant Physiology. 2004;134:1135-45.
cr
1099
Amtmann A, Fischer M, Marsh EL, Stefanovic A, Sanders D, Schachtman DP. The wheat
1105
cDNA LCT1 generates hypersensitivity to sodium in a salt-sensitive yeast strain. Plant
1106
Physiology. 2001;126:1061-71.
us
1104
Ando Y, Nagata S, Yanagisawa S, Yoneyama T. Copper in xylem and phloem saps from
1108
rice (Oryza sativa): the effect of moderate copper concentrations in the growth medium on the
1109
accumulation of five essential metals and a speciation analysis of copper-containing
1110
compounds. Funct Plant Biol. 2013;40:89-100.
M
an
1107
Andres-Colas N, Perea-Garcia A, Puig S, Penaarubia L. Deregulated Copper Transport
1112
Affects Arabidopsis Development Especially in the Absence of Environmental Cycles. Plant
1113
Physiology. 2010;153:170-84.
te
Ac ce p
1114
d
1111
Andres-Colas N, Sancenon V, Rodriguez-Navarro S, Mayo S, Thiele DJ, Ecker JR, et al.
1115
The Arabidopsis heavy metal P-type ATPase HMA5 interacts with metallochaperones and
1116
functions in copper detoxification of roots. Plant Journal. 2006;45:225-36.
1117
Anschutz U, Becker D, Shabala S. Going beyond nutrition: Regulation of potassium
1118
homoeostasis as a common denominator of plant adaptive responses to environment. Journal
1119
of Plant Physiology. 2014;171:670-87.
1120
Aoyama T, Kobayashi T, Takahashi M, Nagasaka S, Usuda K, Kakei Y, et al. OsYSL18 is a
1121
rice iron(III)-deoxymugineic acid transporter specifically expressed in reproductive organs and
1122
phloem of lamina joints. Plant Molecular Biology. 2009;70:681-92.
1123
Apse MP, Blumwald E. Na+ transport in plants. Febs Letters. 2007;581:2247-54. 47 Page 47 of 78
1124
Araki R, Murata J, Murata Y. A Novel Barley Yellow Stripe 1-Like Transporter (HvYSL2)
1125
Localized to the Root Endodermis Transports Metal-Phytosiderophore Complexes. Plant and
1126
Cell Physiology. 2011;52:1931-40. Aranda-Sicilia MN, Cagnac O, Chanroj S, Sze H, Rodriguez-Rosales MP, Venema K.
1128
Arabidopsis KEA2, a homolog of bacterial KefC, encodes a K+/H+ antiporter with a chloroplast
1129
transit peptide. Bba-Biomembranes. 2012;1818:2362-71.
cr
ip t
1127
Ardie SW, Liu S, Takano T. Expression of the AKT1-type K(+) channel gene from
1131
Puccinellia tenuiflora, PutAKT1, enhances salt tolerance in Arabidopsis. Plant Cell Reports.
1132
2010;29:865-74.
us
1130
Arrivault S, Senger T, Kramer U. The Arabidopsis metal tolerance protein AtMTP3
1134
maintains metal homeostasis by mediating Zn exclusion from the shoot under Fe deficiency
1135
and Zn oversupply. Plant Journal. 2006;46:861-79.
M
an
1133
Assunção AGL, Herrero E, Lin YF, Huettel B, Talukdar S, Smaczniak C, et al. Arabidopsis
1137
thaliana transcription factors bZIP19 and bZIP23 regulate the adaptation to zinc deficiency.
1138
Proceedings of the National Academy of Sciences of the United States of America.
1139
2010;107:10296-301.
1141 1142
te
Ac ce p
1140
d
1136
Badri DV, Vivanco JM. Regulation and function of root exudates. Plant Cell and
Environment. 2009;32:666-81.
Barberon M, Zelazny E, Robert S, Conejero G, Curie C, Friml J, et al. Monoubiquitin-
1143
dependent endocytosis of the IRON-REGULATED TRANSPORTER 1 (IRT1) transporter controls
1144
iron uptake in plants. Proceedings of the National Academy of Sciences of the United States of
1145
America. 2011;108:E450-E58.
1146
Barragan V, Leidi EO, Andres Z, Rubio L, De Luca A, Fernandez JA, et al. Ion Exchangers
1147
NHX1 and NHX2 Mediate Active Potassium Uptake into Vacuoles to Regulate Cell Turgor and
1148
Stomatal Function in Arabidopsis. Plant Cell. 2012;24:1127-42.
48 Page 48 of 78
1149 1150
Bashir K, Ishimaru Y, Nishizawa NK. Molecular mechanisms of zinc uptake and translocation in rice. Plant and Soil. 2012;361:189-201. Bashir K, Ishimaru Y, Shimo H, Kakei Y, Senoura T, Takahashi R, et al. Rice phenolics
1152
efflux transporter 2 (PEZ2) plays an important role in solubilizing apoplasmic iron. Soil Sci Plant
1153
Nutr. 2011a;57:803-12.
1155
Bashir K, Ishimaru Y, Shimo H, Nagasaka S, Fujimoto M, Takanashi H, et al. The rice
cr
1154
ip t
1151
mitochondrial iron transporter is essential for plant growth. Nat Commun. 2011b;2.
Bassil E, Ohto MA, Esumi T, Tajima H, Zhu Z, Cagnac O, et al. The Arabidopsis
1157
Intracellular Na+/H+ Antiporters NHX5 and NHX6 Are Endosome Associated and Necessary for
1158
Plant Growth and Development. Plant Cell. 2011;23:224-39.
an
us
1156
Baxter I, Hosmani PS, Rus A, Lahner B, Borevitz JO, Muthukumar B, et al. Root Suberin
1160
Forms an Extracellular Barrier That Affects Water Relations and Mineral Nutrition in
1161
Arabidopsis. Plos Genetics. 2009;5.
d
1163
Benito B, Haro R, Amtmann A, Cuin TA, Dreyer I. The twins K+ and Na+ in plants. Journal of Plant Physiology. 2014;171:723-31.
te
1162
M
1159
Bernal M, Casero D, Singh V, Wilson GT, Grande A, Yang HJ, et al. Transcriptome
1165
Sequencing Identifies SPL7-Regulated Copper Acquisition Genes FRO4/FRO5 and the Copper
1166
Dependence of Iron Homeostasis in Arabidopsis. Plant Cell. 2012;24:738-61.
1167
Ac ce p
1164
Bhatia NP, Walsh KB, Baker AJM. Detection and quantification of ligands involved in
1168
nickel detoxification in a herbaceous Ni hyperaccumulator Stackhousia tryonii Bailey. Journal
1169
of Experimental Botany. 2005;56:1343-49.
1170
Binder BM, Rodriguez FI, Bleecker AB. The Copper Transporter RAN1 Is Essential for
1171
Biogenesis of Ethylene Receptors in Arabidopsis. Journal of Biological Chemistry.
1172
2010;285:37263-70.
49 Page 49 of 78
1173
Cailliatte R, Schikora A, Briat JF, Mari S, Curie C. High-Affinity Manganese Uptake by the
1174
Metal Transporter NRAMP1 Is Essential for Arabidopsis Growth in Low Manganese Conditions.
1175
Plant Cell. 2010;22:904-17. Catty P, Boutigny S, Miras R, Joyard J, Rolland N, Seigneurin-Berny D. Biochemical
1177
Characterization of AtHMA6/PAA1, a Chloroplast Envelope Cu(I)-ATPase. Journal of Biological
1178
Chemistry. 2011;286:36188-97.
cr
ip t
1176
Cellier F, Conejero G, Ricaud L, Luu DT, Lepetit M, Gosti F, et al. Characterization of
1180
AtCHX17, a member of the cation/H+ exchangers, CHX family, from Arabidopsis thaliana
1181
suggests a role in K+ homeostasis. Plant Journal. 2004;39:834-46.
us
1179
Centofanti T, Sayers Z, Cabello-Conejo MI, Kidd P, Nishizawa NK, Kakei Y, et al. Xylem
1183
exudate composition and root-to-shoot nickel translocation in Alyssum species. Plant and Soil.
1184
2013;373:59-75.
1186
M
Chen J, Li LG, Liu ZH, Yuan YJ, Guo LL, Mao DD, et al. Magnesium transporter AtMGT9 is essential for pollen development in Arabidopsis. Cell Research. 2009;19:887-98.
d
1185
an
1182
Cheng NH, Pittman JK, Shigaki T, Lachmansingh J, LeClere S, Lahner B, et al. Functional
1188
association of Arabidopsis CAX1 and CAX3 is required for normal growth and ion homeostasis.
1189
Plant Physiology. 2005;138:2048-60.
Ac ce p
1190
te
1187
Chu HH, Chiecko J, Punshon T, Lanzirotti A, Lahner B, Salt DE, et al. Successful
1191
Reproduction Requires the Function of Arabidopsis YELLOW STRIPE-LIKE1 and YELLOW STRIPE-
1192
LIKE3 Metal-Nicotianamine Transporters in Both Vegetative and Reproductive Structures. Plant
1193
Physiology. 2010;154:197-210.
1194
Conn SJ, Conn V, Tyerman SD, Kaiser BN, Leigh RA, Gilliham M. Magnesium
1195
transporters, MGT2/MRS2-1 and MGT3/MRS2-5, are important for magnesium partitioning
1196
within Arabidopsis thaliana mesophyll vacuoles. New Phytologist. 2011a;190:583-94.
50 Page 50 of 78
1197
Conn SJ, Gilliham M, Athman A, Schreiber AW, Baumann U, Moller I, et al. Cell-Specific
1198
Vacuolar Calcium Storage Mediated by CAX1 Regulates Apoplastic Calcium Concentration, Gas
1199
Exchange, and Plant Productivity in Arabidopsis. Plant Cell. 2011b;23:240-57. Connorton JM, Webster RE, Cheng NH, Pittman JK. Knockout of Multiple Arabidopsis
1201
Cation/H+ Exchangers Suggests Isoform-Specific Roles in Metal Stress Response, Germination
1202
and Seed Mineral Nutrition. Plos One. 2012;7.
cr
1204
Cosio C, DeSantis L, Frey B, Diallo S, Keller C. Distribution of cadmium in leaves of Thiaspi caerulescens. Journal of Experimental Botany. 2005;56:765-75.
us
1203
ip t
1200
Davenport RJ, Munoz-Mayor A, Jha D, Essah PA, Rus A, Tester M. The Na+ transporter
1206
AtHKT1;1 controls retrieval of Na+ from the xylem in Arabidopsis. Plant Cell and Environment.
1207
2007;30:497-507.
1209
Dayod M, Tyerman SD, Leigh RA, Gilliham M. Calcium storage in plants and the
M
1208
an
1205
implications for calcium biofortification. Protoplasma. 2010;247:215-31. Deinlein U, Weber M, Schmidt H, Rensch S, Trampczynska A, Hansen TH, et al. Elevated
1211
Nicotianamine Levels in Arabidopsis halleri Roots Play a Key Role in Zinc Hyperaccumulation.
1212
Plant Cell. 2012;24:708-23.
te
Ac ce p
1213
d
1210
del Pozo T, Cambiazo V, Gonzalez M. Gene expression profiling analysis of copper
1214
homeostasis in Arabidopsis thaliana. Biochemical and Biophysical Research Communications.
1215
2010;393:248-52.
1216
Delhaize E, Gruber BD, Pittman JK, White RG, Leung H, Miao YS, et al. A role for the
1217
AtMTP11 gene of Arabidopsis in manganese transport and tolerance. Plant Journal.
1218
2007;51:198-210.
1219 1220
Demidchik V, Maathuis FJM. Physiological roles of nonselective cation channels in plants: from salt stress to signalling and development. New Phytologist. 2007;175:387-404.
51 Page 51 of 78
1221
Deng W, Luo KM, Li DM, Zheng XL, Wei XY, Smith W, et al. Overexpression of an
1222
Arabidopsis magnesium transport gene, AtMGT1, in Nicotiana benthamiana confers Al
1223
tolerance. Journal of Experimental Botany. 2006;57:4235-43. Drummond RSM, Tutone A, Li YC, Gardner RC. A putative magnesium transporter
1225
AtMRS2-11 is localized to the plant chloroplast envelope membrane system. Plant Science.
1226
2006;170:78-89.
cr
1227
Durrett TP, Gassmann W, Rogers EE. The FRD3-mediated efflux of citrate into the root vasculature is necessary for efficient iron translocation. Plant Physiology. 2007;144:197-205.
us
1228
ip t
1224
Duy D, Stube R, Wanner G, Philippar K. The Chloroplast Permease PIC1 Regulates Plant
1230
Growth and Development by Directing Homeostasis and Transport of Iron. Plant Physiology.
1231
2011;155:1709-22.
an
1229
Edmond C, Shigaki T, Ewert S, Nelson MD, Connorton JM, Chalova V, et al. Comparative
1233
analysis of CAX2-like cation transporters indicates functional and regulatory diversity. Biochem
1234
J. 2009;418:145-54.
d
M
1232
Elbaz B, Shoshani-Knaani N, David-Assael O, Mizrachy-Dagri T, Mizrahi K, Saul H, et al.
1236
High expression in leaves of the zinc hyperaccumulator Arabidopsis halleri of AhMHX, a
1237
homolog of an Arabidopsis thaliana vacuolar metal/proton exchanger. Plant Cell and
1238
Environment. 2006;29:1179-90.
Ac ce p
1239
te
1235
Fedotov PS, Kordel W, Miro M, Peijnenburg WJGM, Wennrich R, Huang PM. Extraction
1240
and Fractionation Methods for Exposure Assessment of Trace Metals, Metalloids, and
1241
Hazardous Organic Compounds in Terrestrial Environments. Crit Rev Env Sci Tec.
1242
2012;42:1117-71.
1243 1244
FNB/IOM. PART III: VITAMINS AND MINERALS.
Essential Guide to Nutrient Requirements: The National Academies Press; 2006. p. 167-422.
1245 1246
Dietary Reference Intakes: The
Forde BG. Glutamate signalling in roots. Journal of Experimental Botany. 2014;65:77987. 52 Page 52 of 78
1247 1248
Fox TC, Guerinot ML. MOLECULAR BIOLOGY OF CATION TRANSPORT IN PLANTS. Annu Rev Plant Phys. 1998;49:669-96. Fukao Y, Ferjani A, Tomioka R, Nagasaki N, Kurata R, Nishimori Y, et al. iTRAQ Analysis
1250
Reveals Mechanisms of Growth Defects Due to Excess Zinc in Arabidopsis. Plant Physiology.
1251
2011;155:1893-907.
ip t
1249
Fulgenzi FR, Peralta ML, Mangano S, Danna CH, Vallejo AJ, Puigdomenech P, et al. The
1253
ionic environment controls the contribution of the barley HvHAK1 transporter to potassium
1254
acquisition. Plant Physiology. 2008;147:252-62.
us
cr
1252
Gajdanowicz P, Michard E, Sandmann M, Rocha M, Correa LGG, Ramirez-Aguilar SJ, et
1256
al. Potassium (K+ ) gradients serve as a mobile energy source in plant vascular tissues.
1257
Proceedings of the National Academy of Sciences of the United States of America.
1258
2011;108:864-69.
M
an
1255
Garcia-Molina A, Andres-Colas N, Perea-Garcia A, del Valle-Tascon S, Penarrubia L, Puig
1260
S. The intracellular Arabidopsis COPT5 transport protein is required for photosynthetic
1261
electron transport under severe copper deficiency. Plant Journal. 2011;65:848-60.
te
d
1259
Gebert M, Meschenmoser K, Svidova S, Weghuber J, Schweyen R, Eifler K, et al. A Root-
1263
Expressed Magnesium Transporter of the MRS2/MGT Gene Family in Arabidopsis thaliana
1264
Allows for Growth in Low-Mg2+ Environments. Plant Cell. 2009;21:4018-30.
1265
Ac ce p
1262
Geiger D, Becker D, Vosloh D, Gambale F, Palme K, Rehers M, et al. Heteromeric
1266
AtKC1.AKT1 Channels in Arabidopsis Roots Facilitate Growth under K+-limiting Conditions.
1267
Journal of Biological Chemistry. 2009;284:21288-95.
1268
Gendre D, Czernic P, Conejero G, Pianelli K, Briat JF, Lebrun M, et al. TcYSL3, a member
1269
of the YSL gene family from the hyper-accumulator Thlaspi caerulescens, encodes a
1270
nicotianamine-Ni/Fe transporter. Plant Journal. 2007;49:1-15.
53 Page 53 of 78
1271
Ghasemi R, Ghaderian SM, Kramer U. Interference of nickel with copper and iron
1272
homeostasis contributes to metal toxicity symptoms in the nickel hyperaccumulator plant
1273
Alyssum inflatum. New Phytologist. 2009;184:566-80. Guo WJ, Meetam M, Goldsbrough PB. Examining the specific contributions of
1275
individual Arabidopsis metallothioneins to copper distribution and metal tolerance. Plant
1276
Physiology. 2008;146:1697-706.
cr
ip t
1274
Gustin JL, Loureiro ME, Kim D, Na G, Tikhonova M, Salt DE. MTP1-dependent Zn
1278
sequestration into shoot vacuoles suggests dual roles in Zn tolerance and accumulation in Zn-
1279
hyperaccumulating plants. Plant Journal. 2009;57:1116-27.
1284 1285 1286 1287 1288
an
M
1283
Haferkamp I, Linka N. Functional expression and characterisation of membrane transport proteins. Plant Biology. 2012;14:675-90.
Hamamoto S, Uozumi N. Organelle-localized potassium transport systems in plants.
d
1282
facilitator family of ion transporters. Bmc Evol Biol. 2011;11.
Journal of Plant Physiology. 2014;171:743-47.
te
1281
Gustin JL, Zanis MJ, Salt DE. Structure and evolution of the plant cation diffusion
Hänsch R, Mendel RR. Physiological functions of mineral micronutrients (Cu, Zn, Mn,
Ac ce p
1280
us
1277
Fe, Ni, Mo, B, Cl). Curr Opin Plant Biol. 2009;12:259-66. Haro R, Banuelos MA, Senn MAE, Barrero-Gil J, Rodriguez-Navarro A. HKT1 mediates
1289
sodium uniport in roots. Pitfalls in the expression of HKT1 in yeast. Plant Physiology.
1290
2005;139:1495-506.
1291 1292
Harris WR, Sammons RD, Grabiak RC. A speciation model of essential trace metal ions
in phloem. J Inorg Biochem. 2012;116:140-50.
1293
Haydon MJ, Kawachi M, Wirtz M, Hillmer S, Hell R, Kramer U. Vacuolar Nicotianamine
1294
Has Critical and Distinct Roles under Iron Deficiency and for Zinc Sequestration in Arabidopsis.
1295
Plant Cell. 2012;24:724-37.
1296
Hedrich R, Marten I. TPC1-SV Channels Gain Shape. Molecular Plant. 2011;4:428-41. 54 Page 54 of 78
1297 1298
Hinsinger P, Bengough AG, Vetterlein D, Young IM. Rhizosphere: biophysics, biogeochemistry and ecological relevance. Plant and Soil. 2009;321:117-52. Hirschi KD. Nutrient Biofortification of Food Crops. Annu Rev Nutr. 2009;29:401-21.
1300
Horie T, Sugawara M, Okada T, Taira K, Kaothien-Nakayama P, Katsuhara M, et al. Rice
1301
sodium-insensitive potassium transporter, OsHAK5, confers increased salt tolerance in tobacco
1302
BY2 cells. Journal of bioscience and bioengineering. 2011;111:346-56.
cr
ip t
1299
Hu YT, Ming F, Chen WW, Yan JY, Xu ZY, Li GX, et al. TcOPT3, a Member of Oligopeptide
1304
Transporters from the Hyperaccumulator Thlaspi caerulescens, Is a Novel Fe/Zn/Cd/Cu
1305
Transporter. Plos One. 2012;7.
us
1303
Inoue H, Kobayashi T, Nozoye T, Takahashi M, Kakei Y, Suzuki K, et al. Rice OsYSL15 Is
1307
an Iron-regulated Iron(III)-Deoxymugineic Acid Transporter Expressed in the Roots and Is
1308
Essential for Iron Uptake in Early Growth of the Seedlings. Journal of Biological Chemistry.
1309
2009;284:3470-79.
M
an
1306
Irtelli B, Petrucci WA, Navari-Izzo F. Nicotianamine and histidine/proline are,
1311
respectively, the most important copper chelators in xylem sap of Brassica carinata under
1312
conditions of copper deficiency and excess. Journal of Experimental Botany. 2009;60:269-77.
te
Ac ce p
1313
d
1310
Ishimaru Y, Kakei Y, Shimo H, Bashir K, Sato Y, Sato Y, et al. A Rice Phenolic Efflux
1314
Transporter Is Essential for Solubilizing Precipitated Apoplasmic Iron in the Plant Stele. Journal
1315
of Biological Chemistry. 2011;286:24649-55.
1316
Ishimaru Y, Masuda H, Bashir K, Inoue H, Tsukamoto T, Takahashi M, et al. Rice metal-
1317
nicotianamine transporter, OsYSL2, is required for the long-distance transport of iron and
1318
manganese. Plant Journal. 2010;62:379-90.
1319
Ishimaru Y, Takahashi R, Bashir K, Shimo H, Senoura T, Sugimoto K, et al. Characterizing
1320
the role of rice NRAMP5 in Manganese, Iron and Cadmium Transport. Scientific Reports.
1321
2012;2.
55 Page 55 of 78
1322
Jabnoune M, Espeout S, Mieulet D, Fizames C, Verdeil JL, Conejero G, et al. Diversity in
1323
Expression Patterns and Functional Properties in the Rice HKT Transporter Family. Plant
1324
Physiology. 2009;150:1955-71.
1327 1328
ip t
1326
Jeanguenin L, Alcon C, Duby G, Boeglin M, Cherel I, Gaillard I, et al. AtKC1 is a general modulator of Arabidopsis inward Shaker channel activity. The Plant Journal. 2011;67:570-82.
Jeong J, Connolly EL. Iron uptake mechanisms in plants: Functions of the FRO family of
cr
1325
ferric reductases. Plant Science. 2009;176:709-14.
Jeong J, Guerinot ML. Biofortified and bioavailable: The gold standard, for plant-based
1330
diets. Proceedings of the National Academy of Sciences of the United States of America.
1331
2008;105:1777-78.
an
us
1329
Johansson I, Wulfetange K, Poree F, Michard E, Gajdanowicz P, Lacombe B, et al.
1333
External K+ modulates the activity of the Arabidopsis potassium channel SKOR via an unusual
1334
mechanism. Plant Journal. 2006;46:269-81.
d
1336
Jones B, Ljung K. Subterranean space exploration: the development of root system architecture. Curr Opin Plant Biol. 2012;15:97-102.
te
1335
M
1332
Jung HI, Gayomba SR, Rutzke MA, Craft E, Kochian LV, Vatamaniuk OK. COPT6 Is a
1338
Plasma Membrane Transporter That Functions in Copper Homeostasis in Arabidopsis and Is a
1339
Novel Target of SQUAMOSA Promoter-binding Protein-like 7. Journal of Biological Chemistry.
1340
2012;287:33252-67.
1341 1342 1343 1344 1345 1346
Ac ce p
1337
Kabata-Pendias A. Soil–plant transfer of trace elements—an environmental issue.
Geoderma. 2004;122:143-49.
Kakei Y, Ishimaru Y, Kobayashi T, Yamakawa T, Nakanishi H, Nishizawa NK. OsYSL16 plays a role in the allocation of iron. Plant Molecular Biology. 2012;79:583-94. Karley AJ, White PJ. Moving cationic minerals to edible tissues: potassium, magnesium, calcium. Curr Opin Plant Biol. 2009;12:291-98.
56 Page 56 of 78
1347
Kawachi M, Kobae Y, Mori H, Tomioka R, Lee Y, Maeshima M. A Mutant Strain
1348
Arabidopsis thaliana that Lacks Vacuolar Membrane Zinc Transporter MTP1 Revealed the
1349
Latent Tolerance to Excessive Zinc. Plant and Cell Physiology. 2009;50:1156-70. Kim CK, Han JS, Lee HS, Oh JY, Shigaki T, Park SH, et al. Expression of an Arabidopsis
1351
CAX2 variant in potato tubers increases calcium levels with no accumulation of manganese.
1352
Plant Cell Reports. 2006a;25:1226-32.
cr
ip t
1350
Kim SA, Punshon T, Lanzirotti A, Li LT, Alonso JM, Ecker JR, et al. Localization of iron in
1354
Arabidopsis seed requires the vacuolar membrane transporter VIT1. Science. 2006b;314:1295-
1355
98.
1357
Kim YY, Choi H, Segami S, Cho HT, Martinoia E, Maeshima M, et al. AtHMA1
an
1356
us
1353
contributes to the detoxification of excess Zn(II) in Arabidopsis. Plant Journal. 2009;58:737-53. Klaumann S, Nickolaus SD, Furst SH, Starck S, Schneider S, Neuhaus HE, et al. The
1359
tonoplast copper transporter COPT5 acts as an exporter and is required for interorgan
1360
allocation of copper in Arabidopsis thaliana. New Phytologist. 2011;192:393-404.
1363
d
te
1362
Kobayashi T, Nishizawa NK. Iron Uptake, Translocation, and Regulation in Higher Plants. Annual Review of Plant Biology. 2012;63:131-52.
Ac ce p
1361
M
1358
Krebs M, Beyhl D, Gorlich E, Al-Rasheid KAS, Marten I, Stierhof YD, et al. Arabidopsis V-
1364
ATPase activity at the tonoplast is required for efficient nutrient storage but not for sodium
1365
accumulation. Proceedings of the National Academy of Sciences of the United States of
1366
America. 2010;107:3251-56.
1367 1368
Kronzucker HJ, Britto DT. Sodium transport in plants: a critical review. New Phytologist.
2011;189:54-81.
1369
Lanquar V, Lelievre F, Bolte S, Hames C, Alcon C, Neumann D, et al. Mobilization of
1370
vacuolar iron by AtNRAMP3 and AtNRAMP4 is essential for seed germination on low iron.
1371
Embo J. 2005;24:4041-51.
57 Page 57 of 78
Lanquar V, Ramos MS, Lelievre F, Barbier-Brygoo H, Krieger-Liszkay A, Kramer U, et al.
1373
Export of Vacuolar Manganese by AtNRAMP3 and AtNRAMP4 Is Required for Optimal
1374
Photosynthesis and Growth under Manganese Deficiency. Plant Physiology. 2010;152:1986-99.
1375
Larbi A, Morales F, Abadia A, Abadia J. Changes in iron and organic acid concentrations
1376
in xylem sap and apoplastic fluid of iron-deficient Beta vulgaris plants in response to iron
1377
resupply. Journal of Plant Physiology. 2010;167:255-60.
cr
ip t
1372
Larras F, Regier N, Planchon S, Pote J, Renaut J, Cosio C. Physiological and proteomic
1379
changes suggest an important role of cell walls in the high tolerance to metals of Elodea
1380
nuttallii. Journal of Hazardous Materials. 2013;263:575-83.
1383 1384
an
1382
Lee S, An G. Over-expression of OsIRT1 leads to increased iron and zinc accumulations in rice. Plant Cell and Environment. 2009;32:408-16.
Lee S, Kim SA, Lee J, Guerinot ML, An G. Zinc Deficiency-Inducible OsZIP8 Encodes a
M
1381
us
1378
Plasma Membrane-Localized Zinc Transporter in Rice. Molecules and Cells. 2010;29:551-58. Lee SM, Kim HS, Han HJ, Moon BC, Kim CY, Harper JF, et al. Identification of a
1386
calmodulin-regulated autoinhibited Ca2+-ATPase (ACA11) that is localized to vacuole
1387
membranes in Arabidopsis. Febs Letters. 2007;581:3943-49.
te
Ac ce p
1388
d
1385
Li XY, Chanroj S, Wu ZY, Romanowsky SM, Harper JF, Sze H. A distinct endosomal
1389
Ca2+/Mn2+ pump affects root growth through the secretory process. Plant Physiology.
1390
2008;147:1675-89.
1391
Lin YF, Liang HM, Yang SY, Boch A, Clemens S, Chen CC, et al. Arabidopsis IRT3 is a zinc-
1392
regulated and plasma membrane localized zinc/iron transporter. New Phytologist.
1393
2009;182:392-404.
1394 1395 1396 1397
Ma W, Smigel A, Verma R, Berkowitz GA. Cyclic nucleotide gated channels and related signaling components in plant innate immunity. Plant Signaling & Behavior. 2009;4:277-82. Maathuis FJM. Physiological functions of mineral macronutrients. Curr Opin Plant Biol. 2009;12:250-58. 58 Page 58 of 78
1398 1399
Manohar M, Shigaki T, Hirschi KD. Plant cation/H+ exchangers (CAXs): biological functions and genetic manipulations. Plant Biology. 2011;13:561-69. Mao DD, Tian LF, Li LG, Chen J, Deng PY, Li DP, et al. AtMGT7: An Arabidopsis Gene
1401
Encoding a Low-Affinity Magnesium Transporter. Journal of Integrative Plant Biology.
1402
2008;50:1530-38.
ip t
1400
Mari S, Gendre D, Pianelli K, Ouerdane L, Lobinski R, Briat JF, et al. Root-to-shoot long-
1404
distance circulation of nicotianamine and nicotianamine-nickel chelates in the metal
1405
hyperaccumulator Thlaspi caerulescens. Journal of Experimental Botany. 2006;57:4111-22.
1408 1409
us
Context. Annual Review of Plant Biology. 2012;63:183-213.
an
1407
Martinoia E, Meyer S, De Angeli A, Nagy R. Vacuolar Transporters in Their Physiological
McAinsh MR, Pittman JK. Shaping the calcium signature. New Phytologist. 2009;181:275-94.
M
1406
cr
1403
Mian A, Oomen RJFJ, Isayenkov S, Sentenac H, Maathuis FJM, Very AA. Over-
1411
expression of an Na+- and K+-permeable HKT transporter in barley improves salt tolerance.
1412
Plant Journal. 2011;68:468-79.
te
d
1410
Mikkelsen MD, Pedas P, Schiller M, Vincze E, Mills RF, Borg S, et al. Barley HvHMA1 Is a
1414
Heavy Metal Pump Involved in Mobilizing Organellar Zn and Cu and Plays a Role in Metal
1415
Loading into Grains. Plos One. 2012;7.
1416
Ac ce p
1413
Millaleo R, Reyes-Diaz M, Alberdi M, Ivanov AG, Krol M, Huner NPA. Excess manganese
1417
differentially inhibits photosystem I versus II in Arabidopsis thaliana. Journal of Experimental
1418
Botany. 2013;64:343-54.
1419
Mills RF, Doherty ML, Lopez-Marques RL, Weimar T, Dupree P, Palmgren MG, et al.
1420
ECA3, a Golgi-localized P-2A-type ATPase, plays a crucial role in manganese nutrition in
1421
Arabidopsis. Plant Physiology. 2008;146:116-28.
1422 1423
Mills RF, Peaston KA, Runions J, Williams LE. HvHMA2, a P-1B-ATPase from Barley, Is Highly Conserved among Cereals and Functions in Zn and Cd Transport. Plos One. 2012;7. 59 Page 59 of 78
1424
Milner MJ, Seamon J, Craft E, Kochian LV. Transport properties of members of the ZIP
1425
family in plants and their role in Zn and Mn homeostasis. Journal of Experimental Botany.
1426
2013;64:369-81. Moller IS, Gilliham M, Jha D, Mayo GM, Roy SJ, Coates JC, et al. Shoot Na+ Exclusion
1428
and Increased Salinity Tolerance Engineered by Cell Type-Specific Alteration of Na+ Transport
1429
in Arabidopsis. Plant Cell. 2009;21:2163-78.
cr
ip t
1427
Morel M, Crouzet J, Gravot A, Auroy P, Leonhardt N, Vavasseur A, et al. AtHMA3, a P-
1431
1B-ATPase Allowing Cd/Zn/Co/Pb Vacuolar Storage in Arabidopsis. Plant Physiology.
1432
2009;149:894-904.
us
1430
Moreno-Risueno MA, Van Norman JM, Moreno A, Zhang JY, Ahnert SE, Benfey PN.
1434
Oscillating Gene Expression Determines Competence for Periodic Arabidopsis Root Branching.
1435
Science. 2010;329:1306-11.
1437
M
Morrissey J, Baxter IR, Lee J, Li LT, Lahner B, Grotz N, et al. The Ferroportin Metal Efflux Proteins Function in Iron and Cobalt Homeostasis in Arabidopsis. Plant Cell. 2009;21:3326-38.
d
1436
an
1433
Mortimer JC, Laohavisit A, Macpherson N, Webb A, Brownlee C, Battey NH, et al.
1439
Annexins: multifunctional components of growth and adaptation. Journal of Experimental
1440
Botany. 2008;59:533-44.
Ac ce p
1441
te
1438
Mukherjee I, Campbell NH, Ash JS, Connolly EL. Expression profiling of the Arabidopsis
1442
ferric chelate reductase (FRO) gene family reveals differential regulation by iron and copper.
1443
Planta. 2006;223:1178-90.
1444
Murata Y, Harada E, Sugase K, Namba K, Horikawa M, Ma JF, et al. Specific transporter
1445
for iron(III)-phytosiderophore complex involved in iron uptake by barley roots. Pure and
1446
Applied Chemistry. 2008;80:2689-97.
1447
Nieves-Cordones M, Aleman F, Martinez V, Rubio F. The Arabidopsis thaliana HAK5 K+
1448
Transporter Is Required for Plant Growth and K+ Acquisition from Low K+ Solutions under
1449
Saline Conditions. Molecular Plant. 2010;3:326-33. 60 Page 60 of 78
1450
Nieves-Cordones M, Aleman F, Martinez V, Rubio F. K+ uptake in plant roots. The
1451
systems involved, their regulation and parallels in other organisms. Journal of Plant Physiology.
1452
2014;171:688-95.
1454
Nishida S, Aisu A, Mizuno T. Induction of IRT1 by the nickel-induced iron-deficient response in Arabidopsis. Plant Signaling & Behavior. 2012;7:329-31.
ip t
1453
Nishiyama R, Kato M, Nagata S, Yanagisawa S, Yoneyama T. Identification of Zn-
1456
Nicotianamine and Fe-2'-Deoxymugineic Acid in the Phloem Sap from Rice Plants (Oryza sativa
1457
L.). Plant and Cell Physiology. 2012;53:381-90.
us
cr
1455
Nozoye T, Nagasaka S, Kobayashi T, Takahashi M, Sato Y, Sato Y, et al.
1459
Phytosiderophore Efflux Transporters Are Crucial for Iron Acquisition in Graminaceous Plants.
1460
Journal of Biological Chemistry. 2011;286:5446-54.
1462
Nozoye T, Nakanishi H, Nishizawa NK. Characterizing the Crucial Components of Iron
M
1461
an
1458
Homeostasis in the Maize Mutants ys1 and ys3. Plos One. 2013;8. Oomen RJFJ, Wu J, Lelievre F, Blanchet S, Richaud P, Barbier-Brygoo H, et al. Functional
1464
characterization of NRAMP3 and NRAMP4 from the metal hyperaccumulator Thlaspi
1465
caerulescens. New Phytologist. 2009;181:637-50.
1467 1468 1469 1470 1471 1472 1473 1474 1475
te
Ac ce p
1466
d
1463
Pal R, Rai JPN. Phytochelatins: Peptides Involved in Heavy Metal Detoxification. Appl
Biochem Biotech. 2010;160:945-63. Palm C, Sanchez P, Ahamed S, Awiti A. Soils: A contemporary perspective. Annu Rev
Env Resour. 2007;32:99-129.
Palmgren MG. Plant plasma membrane H+-ATPases: Powerhouses for nutrient uptake.
Annu Rev Plant Phys. 2001a;52:817-45. Palmgren MG. PLANT PLASMA MEMBRANE H+-ATPases: Powerhouses for Nutrient Uptake. Annu Rev Plant Phys. 2001b;52:817-45. Park S, Kang TS, Kim CK, Han JS, Kim S, Smith RH, et al. Genetic manipulation for enhancing calcium content in potato tuber. J Agr Food Chem. 2005;53:5598-603. 61 Page 61 of 78
1476 1477
Park S, Kim CK, Pike LM, Smith RH, Hirschi KD. Increased calcium in carrots by expression of an Arabidopsis H+/Ca2+ transporter. Mol Breeding. 2004;14:275-82. Pedas P, Ytting CK, Fuglsang AT, Jahn TP, Schjoerring JK, Husted S. Manganese
1479
efficiency in barley: Identification and characterization of the metal ion transporter HvIRT1.
1480
Plant Physiology. 2008;148:455-66.
ip t
1478
Petersson SV, Johansson AI, Kowalczyk M, Makoveychuk A, Wang JY, Moritz T, et al. An
1482
Auxin Gradient and Maximum in the Arabidopsis Root Apex Shown by High-Resolution Cell-
1483
Specific Analysis of IAA Distribution and Synthesis. Plant Cell. 2009;21:1659-68.
1486 1487
us
of beneficial elements. Curr Opin Plant Biol. 2009;12:267-74.
an
1485
Pilon-Smits EAH, Quinn CF, Tapken W, Malagoli M, Schiavon M. Physiological functions
Pilon M, Cohu CM, Ravet K, Abdel-Ghany SE, Gaymard F. Essential transition metal homeostasis in plants. Curr Opin Plant Biol. 2009;12:347-57.
M
1484
cr
1481
Pineau C, Loubet S, Lefoulon C, Chalies C, Fizames C, Lacombe B, et al. Natural
1489
Variation at the FRD3 MATE Transporter Locus Reveals Cross-Talk between Fe Homeostasis
1490
and Zn Tolerance in Arabidopsis thaliana. Plos Genetics. 2012;8.
1492 1493
te
Pinto E, Aguiar AARM, Ferreira IMPLVO. Influence of Soil Chemistry and Plant
Ac ce p
1491
d
1488
Physiology in the Phytoremediation of Cu, Mn, and Zn. Crit Rev Plant Sci. 2014a;33:351-73. Pinto E, Almeida AA, Aguiar AARM, Ferreira IMPLVO. Changes in macrominerals, trace
1494
elements and pigments content during lettuce (Lactuca sativa L.) growth: Influence of soil
1495
composition. Food Chem. 2014b;152:603-11.
1496
Pinto E, Almeida AA, Ferreira IMPLVO. Assessment of metal(loid)s phytoavailability in
1497
intensive agricultural soils by the application of single extractions to rhizosphere soil.
1498
Ecotoxicology and Environmental Safety. 2015;113:418-24.
1499 1500 1501
Pittman JK. Managing the manganese: molecular mechanisms of manganese transport and homeostasis. New Phytologist. 2005;167:733-42. Pittman JK. Vacuolar Ca2+ uptake. Cell Calcium. 2011;50:139-46. 62 Page 62 of 78
1502
Pottosin I, Dobrovinskaya O. Non-selective cation channels in plasma and vacuolar
1503
membranes and their contribution to K+ transport. Journal of Plant Physiology. 2014;171:732-
1504
42.
1506
Puig S, Penarrubia L. Placing metal micronutrients in context: transport and
ip t
1505
distribution in plants. Curr Opin Plant Biol. 2009;12:299-306.
Ranf S, Wunnenberg P, Lee J, Becker D, Dunkel M, Hedrich R, et al. Loss of the vacuolar
1508
cation channel, AtTPC1, does not impair Ca2+ signals induced by abiotic and biotic stresses.
1509
Plant Journal. 2008;53:287-99.
us
cr
1507
Rellan-Alvarez R, Abadia J, Alvarez-Fernandez A. Formation of metal-nicotianamine
1511
complexes as affected by pH, ligand exchange with citrate and metal exchange. A study by
1512
electrospray ionization time-of-flight mass spectrometry. Rapid Commun Mass Sp.
1513
2008;22:1553-62.
M
an
1510
Rellan-Alvarez R, Giner-Martinez-Sierra J, Orduna J, Orera I, Rodriguez-Castrillon JA,
1515
Garcia-Alonso JI, et al. Identification of a Tri-Iron(III), Tri-Citrate Complex in the Xylem Sap of
1516
Iron-Deficient Tomato Resupplied with Iron: New Insights into Plant Iron Long-Distance
1517
Transport. Plant and Cell Physiology. 2010;51:91-102.
te
Ac ce p
1518
d
1514
Richau KH, Kozhevnikova AD, Seregin IV, Vooijs R, Koevoets PLM, Smith JAC, et al.
1519
Chelation by histidine inhibits the vacuolar sequestration of nickel in roots of the
1520
hyperaccumulator Thlaspi caerulescens. New Phytologist. 2009;183:106-16.
1521
Rubio F, Nieves-Cordones M, Aleman F, Martinez V. Relative contribution of AtHAK5
1522
and AtAKT1 to K(+) uptake in the high-affinity range of concentrations. Physiologia Plantarum.
1523
2008;134:598-608.
1524
Saito A, Saito M, Ichikawa Y, Yoshiba M, Tadano T, Miwa E, et al. Difference in the
1525
distribution and speciation of cellular nickel between nickel-tolerant and non-tolerant
1526
Nicotiana tabacum L. cv. BY-2 cells. Plant Cell and Environment. 2010;33:174-87.
63 Page 63 of 78
1527
Sancenon V, Puig S, Mateu-Andres I, Dorcey E, Thiele DJ, Penarrubia L. The Arabidopsis
1528
copper transporter COPT1 functions in root elongation and pollen development. Journal of
1529
Biological Chemistry. 2004;279:15348-55. Sanchez-Pardo B, Fernandez-Pascual M, Zornoza P. Copper microlocalisation,
1531
ultrastructural alterations and antioxidant responses in the nodules of white lupin and soybean
1532
plants grown under conditions of copper excess. Environmental and Experimental Botany.
1533
2012;84:52-60.
1537 1538 1539
cr
us
1536
roots. New Phytologist. 2009;183:1072-84.
Sasaki A, Yamaji N, Xia JX, Ma JF. OsYSL6 Is Involved in the Detoxification of Excess
an
1535
Santi S, Schmidt W. Dissecting iron deficiency-induced proton extrusion in Arabidopsis
Manganese in Rice. Plant Physiology. 2011;157:1832-40.
Sassi A, Mieulet D, Khan I, Moreau B, Gaillard I, Sentenac H, et al. The Rice Monovalent
M
1534
ip t
1530
Cation Transporter OsHKT2;4: Revisited Ionic Selectivity. Plant Physiology. 2012;160:498-510. Schaaf G, Honsbein A, Meda AR, Kirchner S, Wipf D, von Wiren N. AtIREG2 encodes a
1541
tonoplast transport protein involved in iron-dependent nickel detoxification in Arabidopsis
1542
thaliana roots. Journal of Biological Chemistry. 2006;281:25532-40.
te
Ac ce p
1543
d
1540
Seo PJ, Park J, Park MJ, Kim YS, Kim SG, Jung JH, et al. A Golgi-localized MATE
1544
transporter mediates iron homoeostasis under osmotic stress in Arabidopsis. Biochem J.
1545
2012;442:551-61.
1546
Shanmugam V, Lo JC, Wu CL, Wang SL, Lai CC, Connolly EL, et al. Differential expression
1547
and regulation of iron-regulated metal transporters in Arabidopsis halleri and Arabidopsis
1548
thaliana - the role in zinc tolerance. New Phytologist. 2011;190:125-37.
1549 1550 1551 1552
Shaul O. Magnesium transport and function in plants: the tip of the iceberg. Biometals. 2002;15:309-23. Shaul O, Hilgemann DW, de-Almeida-Engler J, Van Montagu M, Inzé D, Galili G. Cloning and characterization of a novel Mg2+/H+ exchanger. The EMBO Journal. 1999;18:3973-80. 64 Page 64 of 78
1553 1554
Shi HZ, Quintero FJ, Pardo JM, Zhu JK. The putative plasma membrane Na+/H+ antiporter SOS1 controls long-distance Na+ transport in plants. Plant Cell. 2002;14:465-77. Siemianowski O, Barabasz A, Kendziorek M, Ruszczynska A, Bulska E, Williams LE, et al.
1556
HMA4 expression in tobacco reduces Cd accumulation due to the induction of the apoplastic
1557
barrier. Journal of Experimental Botany. 2014;65:1125-39.
ip t
1555
Siemianowski O, Mills RF, Williams LE, Antosiewicz DM. Expression of the P-1B-type
1559
ATPase AtHMA4 in tobacco modifies Zn and Cd root to shoot partitioning and metal tolerance.
1560
Plant Biotechnol J. 2011;9:64-74.
us
1562
Sinclair SA, Kramer U. The zinc homeostasis network of land plants. Bba-Mol Cell Res. 2012;1823:1553-67.
an
1561
cr
1558
Song CP, Guo Y, Qiu QS, Lambert G, Galbraith DW, Jagendorf A, et al. A probable
1564
Na+(K+)/H+ exchanger on the chloroplast envelope functions in pH homeostasis and
1565
chloroplast development in Arabidopsis thaliana. Proceedings of the National Academy of
1566
Sciences of the United States of America. 2004;101:10211-16.
d
M
1563
Song WY, Choi KS, Kim DY, Geisler M, Park J, Vincenzetti V, et al. Arabidopsis PCR2 Is a
1568
Zinc Exporter Involved in Both Zinc Extrusion and Long-Distance Zinc Transport. Plant Cell.
1569
2010;22:2237-52.
Ac ce p
1570
te
1567
Stacey MG, Patel A, McClain WE, Mathieu M, Remley M, Rogers EE, et al. The
1571
Arabidopsis AtOPT3 protein functions in metal homeostasis and movement of iron to
1572
developing seeds. Plant Physiology. 2008;146:589-601.
1573
Sunarpi, Horie T, Motoda J, Kubo M, Yang H, Yoda K, et al. Enhanced salt tolerance
1574
mediated by AtHKT1 transporter-induced Na+ unloading from xylem vessels to xylem
1575
parenchyma cells. Plant Journal. 2005;44:928-38.
1576
Suzuki M, Tsukamoto T, Inoue H, Watanabe S, Matsuhashi S, Takahashi M, et al.
1577
Deoxymugineic acid increases Zn translocation in Zn-deficient rice plants. Plant Molecular
1578
Biology. 2008;66:609-17. 65 Page 65 of 78
1579 1580
Swarbreck SM, Colaco R, Davies JM. Plant calcium-permeable channels. Plant Physiology. 2013;163:514-22. Takahashi R, Bashir K, Ishimaru Y, Nishizawa NK, Nakanishi H. The role of heavy-metal
1582
ATPases, HMAs, in zinc and cadmium transport in rice. Plant signaling & behavior.
1583
2012a;7:1605-7.
ip t
1581
Takahashi R, Ishimaru Y, Senoura T, Shimo H, Ishikawa S, Arao T, et al. The OsNRAMP1
1585
iron transporter is involved in Cd accumulation in rice. Journal of Experimental Botany.
1586
2011;62:4843-50.
us
cr
1584
Takahashi R, Ishimaru Y, Shimo H, Ogo Y, Senoura T, Nishizawa NK, et al. The OsHMA2
1588
transporter is involved in root-to-shoot translocation of Zn and Cd in rice. Plant Cell and
1589
Environment. 2012b;35:1948-57.
an
1587
Tanoi K, Saito T, Iwata N, Kobayashi NI, Nakanishi TM. The analysis of magnesium
1591
transport system from external solution to xylem in rice root. Soil Sci Plant Nutr. 2011;57:265-
1592
71.
d
M
1590
Tapken W, Ravet K, Pilon M. Plastocyanin Controls the Stabilization of the Thylakoid
1594
Cu-transporting P-type ATPase PAA2/HMA8 in Response to Low Copper in Arabidopsis. Journal
1595
of Biological Chemistry. 2012;287:18544-50.
Ac ce p
1596
te
1593
Tian SK, Lu LL, Yang XE, Labavitch JM, Huang YY, Brown P. Stem and leaf sequestration
1597
of zinc at the cellular level in the hyperaccumulator Sedum alfredii. New Phytologist.
1598
2009;182:116-26.
1599
Ueno D, Yamaji N, Ma JF. Further characterization of ferric-phytosiderophore
1600
transporters ZmYS1 and HvYS1 in maize and barley. Journal of Experimental Botany.
1601
2009;60:3513-20.
1602
van Kleeff PJM, Jaspert N, Li KW, Rauch S, Oecking C, de Boer AH. Higher order
1603
Arabidopsis 14-3-3 mutants show 14-3-3 involvement in primary root growth both under
1604
control and abiotic stress conditions. Journal of Experimental Botany. 2014;65:5877-88. 66 Page 66 of 78
1605
Vert G, Barberon M, Zelazny E, Seguela M, Briat JF, Curie C. Arabidopsis IRT2
1606
cooperates with the high-affinity iron uptake system to maintain iron homeostasis in root
1607
epidermal cells. Planta. 2009;229:1171-79. Vert G, Grotz N, Dedaldechamp F, Gaymard F, Guerinot ML, Briat JF, et al. IRT1, an
1609
Arabidopsis transporter essential for iron uptake from the soil and for plant growth. Plant Cell.
1610
2002;14:1223-33.
cr
ip t
1608
Very AA, Nieves-Cordones M, Daly M, Khan I, Fizames C, Sentenac H. Molecular biology
1612
of K+ transport across the plant cell membrane: What do we learn from comparison between
1613
plant species? Journal of Plant Physiology. 2014;171:748-69.
us
1611
Voelker C, Gomez-Porras JL, Becker D, Hamamoto S, Uozumi N, Gambale F, et al. Roles
1615
of tandem-pore K plus channels in plants - a puzzle still to be solved. Plant Biology. 2010;12:56-
1616
63.
1618
M
Wang Y, Wu WH. Potassium Transport and Signaling in Higher Plants. Annual Review of Plant Biology. 2013;64:451-76.
d
1617
an
1614
Waters BM, Grusak MA. Whole-plant mineral partitioning throughout the life cycle in
1620
Arabidopsis thaliana ecotypes Columbia, Landsberg erecta, Cape Verde Islands, and the
1621
mutant line ysl1ysl3. New Phytologist. 2008;177:389-405.
Ac ce p
1622
te
1619
White PJ, Bowen HC, Demidchik V, Nichols C, Davies JA. Genes for calcium-permeable
1623
channels in the plasma membrane of plant root cells. Bba-Biomembranes. 2002;1564:299-309.
1624
White PJ, Broadley MR. Calcium in plants. Annals of Botany. 2003;92:487-511.
1625 1626 1627 1628
White PJ, Brown PH. Plant nutrition for sustainable development and global health.
Annals of Botany. 2010;105:1073-80. Wu B, Becker JS. Imaging techniques for elements and element species in plant science. Metallomics. 2012;4:403-16.
67 Page 67 of 78
1629
Xu WF, Jia LG, Baluska F, Ding GC, Shi WM, Ye NH, et al. PIN2 is required for the
1630
adaptation of Arabidopsis roots to alkaline stress by modulating proton secretion. Journal of
1631
Experimental Botany. 2012a;63:6105-14. Xu WF, Jia LG, Shi WM, Baluska F, Kronzucker HJ, Liang JS, et al. The Tomato 14-3-3
1633
Protein TFT4 Modulates H+ Efflux, Basipetal Auxin Transport, and the PKS5-J3 Pathway in the
1634
Root Growth Response to Alkaline Stress. Plant Physiology. 2013a;163:1817-28.
cr
ip t
1632
Xu WF, Jia LG, Shi WM, Liang JS, Zhou F, Li QF, et al. Abscisic acid accumulation
1636
modulates auxin transport in the root tip to enhance proton secretion for maintaining root
1637
growth under moderate water stress. New Phytologist. 2013b;197:139-50.
us
1635
Xu WF, Shi WM, Jia LG, Liang JS, Zhang JH. TFT6 and TFT7, two different members of
1639
tomato 14-3-3 gene family, play distinct roles in plant adaption to low phosphorus stress. Plant
1640
Cell and Environment. 2012b;35:1393-406.
M
an
1638
Yamasaki H, Hayashi M, Fukazawa M, Kobayashi Y, Shikanai T. SQUAMOSA Promoter
1642
Binding Protein-Like7 Is a Central Regulator for Copper Homeostasis in Arabidopsis. Plant Cell.
1643
2009;21:347-61.
te
d
1641
Yang YQ, Qin YX, Xie CG, Zhao FY, Zhao JF, Liu DF, et al. The Arabidopsis Chaperone J3
1645
Regulates the Plasma Membrane H+-ATPase through Interaction with the PKS5 Kinase. Plant
1646
Cell. 2010;22:1313-32.
1647
Ac ce p
1644
Yao X, Horie T, Xue SW, Leung HY, Katsuhara M, Brodsky DE, et al. Differential Sodium
1648
and Potassium Transport Selectivities of the Rice OsHKT2;1 and OsHKT2;2 Transporters in Plant
1649
Cells. Plant Physiology. 2010;152:341-55.
1650
Yao YN, Xu G, Mou DL, Wang JR, Ma JB. Subcellular Mn compartation, anatomic and
1651
biochemical changes of two grape varieties in response to excess manganese. Chemosphere.
1652
2012;89:150-57.
1653 1654
Yokosho K, Yamaji N, Ueno D, Mitani N, Ma JF. OsFRDL1 Is a Citrate Transporter Required for Efficient Translocation of Iron in Rice. Plant Physiology. 2009;149:297-305. 68 Page 68 of 78
1655 1656
Yuan M, Li X, Xiao J, Wang S. Molecular and functional analyses of COPT/Ctr-type copper transporter-like gene family in rice. BMC Plant Biology. 2011;11:69.
1657
plants. Plant and Soil. 2010;326:45-60.
ip t
1658
Zhang JL, Flowers TJ, Wang SM. Mechanisms of sodium uptake by roots of higher
Zhang JL, Flowers TJ, Wang SM. Differentiation of low-affinity Na+ uptake pathways
1660
and kinetics of the effects of K+ on Na+ uptake in the halophyte Suaeda maritima. Plant and
1661
Soil. 2013;368:629-40.
cr
1659
Zhang WJ, To JPC, Cheng CY, Schaller GE, Kieber JJ. Type-A response regulators are
1663
required for proper root apical meristem function through post-transcriptional regulation of
1664
PIN auxin efflux carriers. Plant Journal. 2011;68:1-10.
an
us
1662
Zhang Y, Xu YH, Yi HY, Gong JM. Vacuolar membrane transporters OsVIT1 and OsVIT2
1666
modulate iron translocation between flag leaves and seeds in rice. Plant Journal. 2012;72:400-
1667
10.
M
1665
Zhao HJ, Wu LQ, Chai TY, Zhang YX, Tan JJ, Ma SW. The effects of copper, manganese
1669
and zinc on plant growth and elemental accumulation in the manganese-hyperaccumulator
1670
Phytolacca americana. Journal of Plant Physiology. 2012;169:1243-52.
1672 1673
te
Ac ce p
1671
d
1668
Zhao J, Cheng NH, Motes CM, Blancaflor EB, Moore M, Gonzales N, et al. AtCHX13 is a
plasma membrane K+ transporter. Plant Physiology. 2008;148:796-807. Zheng LQ, Yamaji N, Yokosho K, Ma JF. YSL16 Is a Phloem-Localized Transporter of the
1674
Copper-Nicotianamine Complex That Is Responsible for Copper Distribution in Rice. Plant Cell.
1675
2012;24:3767-82.
1676 1677
Zörb C, Senbayram M, Peiter E. Potassium in agriculture – Status and perspectives. Journal of Plant Physiology. 2014;171:656-69.
1678
Zorrig W, Abdelly C, Berthomieu P. The phylogenetic tree gathering the plant
1679
Zn/Cd/Pb/Co P1B-ATPases appears to be structured according to the botanical families.
1680
Comptes rendus biologies. 2011;334:863-71.
1681
69 Page 69 of 78
Table 1. Abbreviations list Ca2+-ATPase
BCD
Bush-and-chlorotic-dwarf
CAX
Ca2+/H+ exchanger
CDF
Cation diffusion facilitator
CEC
Cation exchange capacity
CHX
Cation/H+ exchanger
CNGC
Cyclic-nucleotide gated channel
COPT
Copper transporter
DA-NSCC
Depolarization-activated nonselective cation channel
DMA
Deoxymugineic acid
ECA
Endomembrane-type Ca2+-ATPase
ENA
Efflux transporter of nicotianamine
FRD
Ferric reductase defective
FRDL
Ferric reductase defective-like
FRO
Ferric-chelate reductase oxidase
GLR
Ionotropic glutamate receptor
GSH
Glutathione
HA
H+-ATPase
HA-NSCC
Hyperpolarization-activated nonselective cation channel
HAK/KT/KUP
K+/H+ symporters
HATS
High-affinity transporter system
HKT/Trk
K+/Na+ symporters
HMA
Heavy metal associated
IREG/FPN
Iron-regulated/Ferroportin
IRT
Iron-regulated transporter
KCO
Kir-like K+ channel
KEA
K+ efflux antiporter
LATS
Low-affinity transporter system
LCT
Low-affinity cation transporter
MA
Mugineic acid
MATE
Multidrug and toxic compound extrusion
MRS2/MGT
Magnesium transporter
te
d
M
an
us
cr
ip t
ACA
Ac ce p
1681
70 Page 70 of 78
MIT
Mitochondrial transporter
MT
Metallothionein
MTP
Metal tolerance proteins
NA
Nicotianamine
NHX
Na+/H+ antiporter
NRAMP
Natural resistance associated macrophage protein
NSCC
Nonselective cation channel
OM
Organic matter
OPT
Oligopeptide transporter
PC
Phytochelatin
PCR
Plant cadmium resistance
PEZ
Phenolics efflux zero
PIC
Permease in chloroplast
PS
Phytosiderophore
RDA
Recommended dietary allowances
TPC
Two-pore channel
TPK
Two-pore K+ channel
TOM
Transporter of mugineic acid family phytosiderophores
VI-NSCC
Voltage-insensitive nonselective cation channel
VIT
Vacuolar iron transporter
YS
Yellow stripe
YSL
Yellow stripe-Like
ZIF
Zinc-induced facilitator
ZIP
(ZRT-IRT)-like protein
te
d
M
an
us
cr
ip t
Mg2+/H+ antiporter
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