Zinc in plants

Authors

  • Martin R. Broadley,

    1. Plant Sciences Division, School of Biosciences, University of Nottingham, Sutton Bonington Campus, Loughborough LE12 5RD, UK;
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  • Philip J. White,

    1. The Scottish Crop Research Institute, Invergowrie, Dundee DD2 5DA, UK;
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  • John P. Hammond,

    1. Warwick HRI, University of Warwick, Wellesbourne, Warwick CV35 9EF, UK;
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  • Ivan Zelko,

    1. Institute of Chemistry, Slovak Academy of Sciences, Dúbravská cesta 9, SK 84538 Bratislava, Slovakia;
    2. Department of Plant Physiology, Faculty of Natural Sciences, Comenius University, Mlynská dolina B2, SK 84215 Bratislava, Slovakia
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  • Alexander Lux

    1. Institute of Chemistry, Slovak Academy of Sciences, Dúbravská cesta 9, SK 84538 Bratislava, Slovakia;
    2. Department of Plant Physiology, Faculty of Natural Sciences, Comenius University, Mlynská dolina B2, SK 84215 Bratislava, Slovakia
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Author for correspondence: Martin R. Broadley Tel: +44 (0)1159516382 Fax: +44 (0)1159516334 Email: martin.broadley@nottingham.ac.uk

Abstract

Contents

  • Summary 677

  • I. Physical and chemical properties of zinc 678
  • II. Biochemical properties of zinc 678
  • III. Proteins interacting with zinc 678
  • IV. Zinc fluxes in the soil–root–shoot continuum 679
  • V. Zinc in plants 684
  • VI. Plant responses to elevated soil Zn 686
  • Acknowledgements 695

  • References 696

Summary

Zinc (Zn) is an essential component of thousands of proteins in plants, although it is toxic in excess. In this review, the dominant fluxes of Zn in the soil–root–shoot continuum are described, including Zn inputs to soils, the plant availability of soluble Zn2+ at the root surface, and plant uptake and accumulation of Zn. Knowledge of these fluxes can inform agronomic and genetic strategies to address the widespread problem of Zn-limited crop growth. Substantial within-species genetic variation in Zn composition is being used to alleviate human dietary Zn deficiencies through biofortification. Intriguingly, a meta-analysis of data from an extensive literature survey indicates that a small proportion of the genetic variation in shoot Zn concentration can be attributed to evolutionary processes whose effects manifest above the family level. Remarkable insights into the evolutionary potential of plants to respond to elevated soil Zn have recently been made through detailed anatomical, physiological, chemical, genetic and molecular characterizations of the brassicaceous Zn hyperaccumulators Thlaspi caerulescens and Arabidopsis halleri.

I. Physical and chemical properties of zinc

Zinc is a transition metal of atomic number 30 and is the 23rd most abundant element on earth. Zinc has five stable isotopes: 64Zn (48.63%), 66Zn (27.90%), 67Zn (4.90%), 68Zn (18.75%) and 70Zn (0.62%). Heavy and light isotopic enrichments of root and shoot Zn fractions, respectively, have been reported in plants (Weiss et al., 2005). Approximately 30 short-lived Zn radioisotopes occur in the atomic mass range 54–83, and the longest-lived (65Zn, t1/2 = 244.26 d) is frequently used as a Zn tracer in plants. In solution, Zn exists in the +2 oxidation state and, unlike Fe2+ and Cu2+, is redox-stable under physiological conditions as a result of a complete d-shell of electrons (Barak & Helmke, 1993; Auld, 2001). Additionally, Zn2+ has pronounced Lewis acid characteristics because of its small radius to charge ratio (i.e. 0.83 Å, coordination number, CN = 6) compared, for example, with Ca2+ (1.08 Å, CN = 6), and thus forms strong covalent bonds with S, N and O donors. This electron configuration of aqueous Zn2+ complexes favours octahedral coordination geometries (CN = 6), although CN = 4 and CN = 5 geometries also occur (Barak & Helmke, 1993). Zinc forms numerous soluble salts, including halides, sulphates, nitrates, formates, acetates, thiocyanates, perchlorates, fluosilicates, cyanides, alkali metal zincates and Zn-ammonia salts; sparingly soluble compounds, including Zn-ammonium phosphate, Zn hydroxide and Zn carbonate; and a range of soluble and insoluble organic complexes (Lindsay, 1979; Barak & Helmke, 1993).

II. Biochemical properties of zinc

Zinc is typically the second most abundant transition metal in organisms after iron (Fe), and the only metal represented in all six enzyme classes (Enzyme Commission number, EC 1–6; (oxidoreductases, transferases, hydrolases, lyases, isomerases, ligases; http://www.chem.qmul.ac.uk/iubmb/enzyme/; Webb, 1992). Enzymatic function and reactivity are determined by the geometric and binding characteristics of Zn2+-ligand complexes and three primary Zn2+-ligand binding sites are recognized: structural, catalytic and cocatalytic (Auld, 2001; Maret, 2005). Structural Zn sites, in which Zn ensures appropriate protein folding (e.g. alcohol dehydrogenases, protein kinases), comprise four ligands, frequently cysteine (Cys), and no bound water molecule. In catalytic sites, Zn is directly involved in the catalytic function of the enzyme (e.g. carbonic anhydrases); histidine (His) is the primary amino acid and Zn2+ is complexed with water and any three S, N or O donors. In cocatalytic sites, Zn2+ can be used for catalytic, regulatory and structural functions (e.g. superoxide dismutases, purple acid phosphatases, metallo-β-lactamases). At such sites, two or three Zn2+ occur in close proximity and are bridged by amino acid residues, principally aspartic acid (Asp) or glutamic acid (Glu), but also His and potentially a water molecule, but not Cys. A fourth type of Zn2+-ligand binding or protein interface site can behave as a catalytic or structural site. These occur when ligands from the surface of two protein molecules bind to a single Zn atom (e.g. nitric oxide synthases). Zinc binding sites also occur in a wide range of other proteins, membrane lipids and DNA/RNA molecules. The largest class of Zn-binding proteins in organisms is the zinc finger domain containing proteins, which can regulate transcription directly through effects on DNA/RNA binding, and also through site-specific modifications, regulation of chromatin structure, RNA metabolism and protein–protein interactions (Klug, 1999; Englbrecht et al., 2004).

III. Proteins interacting with zinc

In Escherichia coli, femtomolar (1 × 10−15 m) cytosolic concentrations of free Zn2+ induce the activity of Zn influx (Zur) and efflux (ZntR) proteins (Outten & O’Halloran, 2001). Since these concentrations are 106 times lower than one Zn2+ ion per cell, contrasting with a minimal Zn content of an E. coli cell in the millimolar range (c. 200 000 atoms per cell), cytosolic free Zn2+ pools are not thought to persist. Ionic Zn is likely to be excluded from the cytosol via direct transfer between proteins, with > 10% of cellular Zn thought to be tightly bound to just six proteins, including an RNA polymerase expressed at 5000 copies per cell with two Zn atoms bound per copy, and five tRNA synthetases expressed at 2000–3000 copies per cell with a single Zn atom bound per copy (Outten & O’Halloran, 2001). Since there are > 30 further proteins of unknown copy number which require tightly bound Zn (Katayama et al., 2002), and many other proteins, amino acids and nucleotides with lower affinities for Zn, a large Zn-binding overcapacity in the cytosol of E. coli has been predicted (Outten & O’Halloran, 2001). In plant cells, high Zn-status leaf epidermal cell vacuoles, cell walls and cytoplasm (i.e. cytosol and organelles excluding the vacuole) can contain, respectively, 74 305, 11 577 and 3205 µg Zn g−1 DW (dry weight); lower Zn-status leaf mesophyll cell vacuoles, cell walls and cytoplasm contain, respectively, 327, 9353 and ≤ 262 µg Zn g−1 DW; and root cortical vacuoles, cell walls and cytoplasm contain, respectively, ≤ 262, 589 and ≤ 262 µg Zn g−1 DW (Frey et al., 2000). Thus, 9.6 × 1010, 2.7 × 109 and 1.8 × 109 atoms of Zn can occur in leaf epidermal cell vacuoles, cell walls and cytoplasm, respectively. Leaf mesophyll cell vacuoles, cell walls and cytoplasm contain 4.2 × 108, 2.2 × 109 and ≤ 1.5 × 108 atoms of Zn respectively, and root cortical cell vacuoles, cell walls and cytoplasm contain ≤ 3.4 × 108, 1.4 × 108 and ≤ 1.5 × 108 atoms of Zn, respectively. These calculations assume that a plant cell comprises 1 × 10−15 m3 with a mass of 1 × 10−9 g FW (fresh weight); the vacuole, cell wall and cytoplasm occupy 70, 5 and 25%, respectively, of the cell volume; and FW : DW ratios are five, two and four for vacuoles, cell wall and cytoplasm, respectively (Flowers & Yeo, 1992; Frey et al., 2000). It is not yet known what proportion of plant cytoplasmic Zn is present as free Zn2+ or as Zn bound to protein, amino acid, nucleotide or lipid ligands at lower affinities, or compartmentalized into organelles. However, by analogy with E. coli, it seems likely that (i) cytosolic Zn2+ concentration ([Zn2+]cyt) will be vanishingly low to prevent interference with metalloregulatory and other signalling proteins; and (ii) low [Zn2+]cyt will be maintained through high-affinity binding of Zn in the cytosol and through compartmentalization of Zn into cytoplasmic organelles. In mammals, cytoplasmic Zn can be sequestered into vesicles (‘zincosomes’; Beyersmann & Haase, 2001), and this process might also occur in plants.

In the human genome, annotation- and Zn-binding domain-based searches reveal that c. 10% of proteins (i.e. 2800) potentially bind Zn (Andreini et al., 2006), with hundreds more involved in Zn transport and trafficking (Beyersmann & Haase, 2001). A similar in silico study of the complement of Zn-binding proteins in Arabidopsis thaliana (L.) Heynh. was undertaken here. First, protein domains with observed or predicted capabilities for binding Zn were identified from the Pfam database (http://www.sanger.ac.uk/Software/Pfam/; 25 July 2006). Approximately 120 putative Zn-binding protein domains were identified, with 2042 A. thaliana proteins (TAIR6) containing one or more of these domains (Supplementary material, Table S1). Secondly, annotation searches using the words ‘zinc’ or ‘Zn’, corrected for false positives, revealed 1245 genes (Table S2), of which 1096 were common to the domain-based search. Finally, proteins implicated in Zn homeostasis were hand-compiled (Table S3). These included the remaining carbonic anhydrases, alcohol dehydrogenases and proteins with putative Zn transport functions, including PIB-type ATPases, divalent cation transporters and nonspecific cation channels. This third list contained 1635 proteins, 176 of which were unique. In total, 2367 proteins (Table S4) in 181 gene families (Table S5) were identified as Zn-related. Briefly, one or more Gene Ontology molecular function (GO:3674) subcategories were assigned to each gene using GeneSpring GX (Agilent Technologies Inc., Palo Alto, CA, USA) (Table 1). The largest group of Zn-binding proteins in A. thaliana are Zn finger domains, assigned to transcription regulator activity (GO:30528) and binding (GO:5488) functional subcategories. The catalytic activity (GO:3824) subcategory comprises numerous proteins, including those with hydrolase activity (GO:16787, e.g. P1B-ATPases) and transferase activity (GO:16740, e.g. mitogen-activated protein kinases (MAPKs)). The transporter activity (GO:5215) subcategory includes ABC transporters, PIB-ATPases, various divalent cation transporters (for example, the cation diffusion facilitator family (CDFs)), Zn-Fe permeases (ZIPs) and nonspecific cation channels.

Table 1.  Gene Ontology molecular function (GO:3674) assigned to 2367 Zn-related genes in Arabidopsis thaliana, identified through annotation-, domain- and literature-based searches using GeneSpring GX (Agilent Technologies, Inc., Palo Alto, CA, USA) (primary data in Supplementary material, Table S4)
Gene Ontology: molecular function (GO:3674) subcategoriesExample gene familiesNumber of genes with potential role in Zn homeostasisSelected references
Binding (GO:5488)Zinc finger proteins1503Berg & Shi (1996), Klug (1999), Auld (2001)
Squamosa promoter binding proteins Yamasaki et al. (2004)
Metallothioneins Blindauer & Sadler (2005), Roosens et al. (2005), Zimeri et al. (2005)
WRKY family transcription factors Eulgem et al. (2000)
Catalytic activity (GO:3824)Alcohol dehydrogenases 634Chase (1999), Kim et al. (2004)
Carbonic anhydrases Moroney et al. (2001), Tiwari et al. (2005)
Superoxide dismutases Mittler et al. (2004)
Glutathione transferases Edwards & Dixon (2005)
Metallo β-lactamases Auld (2001)
Purple acid phosphatases Olczak et al. (2003)
Mitogen-activated protein kinases (MAPK) Morris (2001)
SET-domain transcriptional regulators Cheng et al. (2005)
PIB-ATPases Williams & Mills (2005)
Transcription regulator activity (GO:30528)Zinc finger proteins (e.g. CCHC, CONSTANS B-Box, Cys2/His2, Dof, GATA) 379Clay & Nelson (2005), Robson et al. (2001), Reyes et al. (2004), Sakamoto et al. (2004), Yanagisawa (2004)
Squamosa promoter binding proteins Yamasaki et al. (2004)
WRKY family transcription factors Ülker & Somssich (2004), Zhang & Wang (2005)
Transporter activity (GO:5215)ABC transporters 254Hall & Williams (2003), Hantke (2005)
PIB-ATPases Williams & Mills (2005)
Divalent cation transporters from several families (e.g. cation diffusion facilitators, CDFs, Zn-Fe permeases, ZIPs) Colangelo & Guerinot (2006), Mäser et al. (2001)
Nonspecific cation channels White et al. (2002a)
Molecular function unknown (GO:5554)  241 
Signal transducer activity (GO:4871)Mitogen-activated protein kinases (MAPK)  26Morris (2001)
Structural molecule activity (GO:5198)40S ribosomal protein S27  12McIntosh & Bonham-Smith (2006)
Translation regulator activity (GO:45182)Translation initiation factors  10Browning (2004)
Enzyme regulator activity (GO:30234)GTPase activator proteins of Rab-related small GTPases-like protein   7Saito et al. (2002)

IV. Zinc fluxes in the soil–root–shoot continuum

1. Zinc inputs to soils

The primary input of Zn to soils is from the chemical and physical weathering of parent rocks. The lithosphere typically comprises 70–80 µg Zn g−1, whilst sedimentary rocks contain 10–120 µg Zn g−1 (Friedland, 1990; Barak & Helmke, 1993; Alloway, 1995). Mean soil Zn concentrations ([Zn]soil) of 50 and 66 µg total Zn g−1 soil are typical for mineral and organic soils, respectively, with most agricultural soils containing 10–300 µg Zn g−1 (Alloway, 1995; Barber, 1995). Zinc occurs in rock-forming minerals as a result of the nonspecific replacement of Mg and Fe with Zn (Barak & Helmke, 1993). Rocks containing weathered Zn minerals, including Zn sulphide (sphalerite, wurtzite), sulphate (zincosite, goslarite), oxide (zincinte, franklinite, gahnite), carbonate (smithsonite), phosphate (hopeite) and silicate (hemimorphite, willemite) minerals, can form ‘calamine’ soils containing extremely high concentrations of Zn and other metals (Barak & Helmke, 1993). For example, in Plombières in Belgium, [Zn]soil exceeds 100 000 µg Zn g−1 (Cappuyns et al., 2006). Such sites are usually localized to a few hectares, although adjacent soils can also have high [Zn]soil through water seepage from ore bodies (Chaney, 1993). Secondary natural inputs of Zn to soils arise because of atmospheric (e.g. volcanoes, forest fires, and surface dusts) and biotic (e.g. decomposition, leaching/washoff from leaf surfaces) processes (Friedland, 1990).

Humans have long influenced Zn inputs to soils. Two thousand years ago, approx. 10 000 tonnes Zn yr−1 were emitted as a result of mining and smelting activities (Nriagu, 1996). Since 1850, emissions have increased 10-fold, peaking at 3.4 Mt Zn yr−1 in the early 1980s, and then declining to 2.7 Mt Zn yr−1 by the early 1990s (Nriagu, 1996). Arctic troposphere Zn concentrations (c. 2 ng Zn m−3 in winter months) are yet to reflect this decline (Gong & Barrie, 2005). The ratio of Zn emissions arising from anthropogenic and natural inputs is estimated to be > 20 : 1 (Friedland, 1990). Other anthropogenic inputs of Zn to soils include fossil fuel combustion, mine waste, phosphatic fertilizers (typically 50–1450 µg Zn g−1), limestone (10–450 µg Zn g−1), manure (15–250 µg Zn g−1), sewage sludge (91–49 000 µg Zn g−1), other agrochemicals, particles from galvanized (Zn-plated) surfaces and rubber mulches (Chaney, 1993; Alloway, 1995). Crop Zn toxicity can occur in Zn-contaminated soils (discussed in Section VI.1).

2. Zinc behaviour in soils

Soil Zn occurs in three primary fractions: (i) water-soluble Zn (including Zn2+ and soluble organic fractions); (ii) adsorbed and exchangeable Zn in the colloidal fraction (associated with clay particles, humic compounds and Al and Fe hydroxides); and (iii) insoluble Zn complexes and minerals (reviewed by Lindsay, 1979; Barrow, 1993; Alloway, 1995; Barber, 1995). The distribution of Zn between soil fractions is determined by soil-specific precipitation, complexation and adsorption reactions. The dominant factor determining soil Zn distribution is pH; Zn is more readily adsorbed on cation exchange sites at higher pH and adsorbed Zn is more readily displaced by CaCl2 at lower pH. Thus, soluble Zn and the ratio of Zn2+ to organic Zn-ligand complexes increase at low pH, especially in soils of low soluble organic matter content. Soil type, soil moisture, mineral and clay types and contents, diffusion and mass flow rates, weathering rates, soil organic matter, soil biota and plant uptake will also affect Zn distribution. Insoluble Zn comprises > 90% of soil Zn and is unavailable for plant uptake. Exchangeable Zn typically ranges from 0.1 to 2 µg Zn g−1. Concentrations of water-soluble Zn in the bulk soil solution ([Zn]bss) are low, typically between 4 × 10−10 and 4 × 10−6 m (Barber, 1995), even in Zn-contaminated soils (Knight et al., 1997). Numerous Zn-ligand complexes can exist in solution which can be difficult to measure directly, and speciation models, based on total dissolved concentrations of elements and ligands, their stability constants and mineral equilibria reactions, are often used to infer [Zn2+]bss (Barak & Helmke, 1993; Zhang & Young, 2006). Zn2+ typically accounts for up to 50% of the soluble Zn fraction and is the dominant plant-available Zn fraction. However, in calcareous soils, Zn2+ may be as low as 10−11−10−9 m and can limit crop growth (Hacisalihoglu & Kochian, 2003; discussed in Section V.1).

Soil Zn fractions in the solid phase can be quantified using sequential extractions or isotopic dilution techniques (Young et al., 2006). For example, Zn extracted by H2O (‘water-soluble’), KNO3 (‘exchangeable’), Na4P2O7 (‘organically bound’), EDTA (‘carbonate/noncrystalline iron occluded’), NH2OH (‘manganese oxide occluded’), Na2S2O4 (‘crystalline iron oxide occluded’), HNO3 (‘sulphides’), HNO3 + H2O2 (‘residual’) represented 0.2, 10.0, 32.5, 7.9, 7.2, 7.5, 3.3 and 32.3% of total soil Zn, respectively, in a mineral soil from Indiana, in the USA (Miller & McFee, 1983). Despite recent methodological advances in measuring metal species in the soil solution (Zhang & Young, 2006), Zn availability at the soil–root interface ([Zn]ext) can still be difficult to determine satisfactorily, especially if Zn uptake by roots is high and Zn-depletion zones develop around the root. Thus, modelling approaches to determine soil and plant effects on Zn dynamics have been developed (Barber & Claassen, 1977; Bar-Yosef et al., 1980; Barber, 1995; Whiting et al., 2003; Qian et al., 2005; Lehto et al., 2006). For example, Whiting et al. (2003) adapted a transport model of Baldwin et al. (1973) to calculate [Zn]ext, using empirically derived soil parameters (Barber, 1995) and empirically derived or inferred plant parameters. The parameters were [Zn]bss (concentration of Zn in the bulk soil solution), D (the effective diffusion coefficient of Zn in the soil solution), b (the Zn-buffering power of the soil), x (the radius of the soil cylinder that can be exploited by the root), α (the root absorption power of plant for Zn), õ (the water flux from the soil to the root surface) and a (the root radius). Values for [Zn]bss vary widely, as described previously, typically in the range 1 × 10−8 to 1 × 10−6m. Values of D also vary widely, increasing at high [Zn]soil, high soil bulk density (air-filled pores increase diffusion resistance), high soil water content and low pH, typically within the range 10−10−10−8 cm2 s−1 for Zn (Barber, 1995). Values of b represent the distribution of Zn between the solution and the solid phases and are estimated from empirically derived Zn adsorption isotherms (Barber, 1995). Values of b are highest at low [Zn]bss, and thus at high cation exchange capacity (CEC) and pH. Values of b typically range from 2.4 to 571 (Barber, 1995), and for vertisols from 217 to 790 (Dang et al., 1994). Paramaters õ and α represent plant-specific transpiration and Zn uptake rates and root morphology. For plants of the same size and with the same root Zn and water-absorbing power, the primary drivers of [Zn]ext are [Zn]bss and b (Whiting et al., 2003). At high b (> 200), [Zn]ext is proportional to, or approximates, [Zn]bss, that is, plant-available Zn is determined by the capacity of the soil to replenish the soluble Zn fraction as it is removed by the plant. However, when b < 200, [Zn]ext is << [Zn]bss, and especially so when b < 50.

Modelling [Zn]ext requires assumptions to minimize complexity and to compensate for a lack of adequate input data. For example, Whiting et al. (2003) assumed that: (i) [Zn]ext was in steady state with roots nonrandomly dispersed at constant density in a finite volume of soil; (ii) only radial transport of Zn occurred; (iii) all Zn in the soil solution was available for uptake; and (iv) no compounds mobilizing Zn from the solid phase were secreted from roots, mycorrhiza or other soil-dwelling organisms. However, models to predict [Zn]ext are continually being improved for both soil (Lehto et al., 2006) and plant (Qian et al., 2005) parameters, and are likely to become invaluable in improving our understanding of whole-plant physiological processes, optimizing crop Zn nutrition through the application of fertilizers and soil amendments, and improving risk assessments for metal-contaminated environments.

3. Zinc fluxes into plants

Zinc is acquired from the soil solution primarily as Zn2+, but also potentially complexed with organic ligands, by roots which feed the shoots via the xylem. The relationship between Zn influx (and uptake or accumulation) to excised roots and intact plants, V, and [Zn]ext is often characterized by the sum of one or more Michaelis–Menten functions, each defined by a Vmax (the rate at [Zn]ext = ∞) and an affinity constant, Km ([Zn]ext when V = 0.5Vmax), plus a linear term, k (V/[Zn]ext). Most detailed kinetic studies report a Michaelis–Menten function with a Km of 1.5–50 µm (with Vmax values of up to 5.74 µmol g−1 FW h−1), and, occasionally, additional Michaelis–Menten functions with higher Km values (Table 2). These Km values are generally higher than [Zn2+]bss (10−11−10−6 m; see Section IV.2). Two studies have reported Michaelis–Menten functions for wheat plants with Km values of 0.6–2.3 nm for Zn2+ uptake (Wheal & Rengel, 1997; Hacisalihoglu et al., 2001). Whilst Km and Vmax do not differ greatly between Zn-efficient and Zn-inefficient wheat varieties, suggesting that the kinetics of Zn influx per se do not play a significant role in Zn efficiency in wheat (Hart et al., 1998, 2002; Hacisalihoglu et al., 2001; Hacisalihoglu & Kochian, 2003), differences in these parameters between Zn-efficient and Zn-inefficient genotypes of sugarcane, rice and tomato have been reported (Bowen, 1973, 1986, 1987). In general, Zn-deficient plants have higher Vmax and comparable Km values to Zn-replete plants (Rengel & Wheal, 1997; Hacisalihoglu et al., 2001). Several authors have attributed a linear term, which is not removed by washing the roots in solutions containing excess cations, to the accumulation of Zn bound strongly to cell walls (Lasat et al., 1996; Hart et al., 1998, 2002; Hacisalihoglu et al., 2001). There has been some discourse about the consequences of Zn binding to cell walls in experiments to determine kinetic parameters for Zn influx to roots (Reid et al., 1996). In the giant alga, Chara, cell walls can be removed and apoplastic effects on Zn influx can be determined directly (Reid et al., 1996). Using this technique, Zn influx to the cytoplasm was described by the sum of a Michaelis–Menten function with a low Km (< 0.1 µm) plus a linear term for [Zn]ext up to 50 µm. These data suggest that most experiments to determine the kinetic parameters of Zn influx to roots may have overestimated Km values and distorted the linear term.

Table 2.  Published Michaelis–Menten functions, describing the relationship between zinc influx (and uptake or accumulation) and excised roots and intact plants, V, according to the terms Vmax (V, when [Zn]ext = ∞), Km ([Zn]ext, when V = 0.5Vmax), and a linear term k (V/[Zn]ext)
Reference[Zn]extm)Function 1Function 2Function 3kNotesb
Kmm)Vmaxa (nmol g−1 h−1)Kmm)Vmaxa (nmol g−1 h−1)Kmm)Vmaxa (nmol g−1 h−1)
  • a

    Vmax data expressed on a root fresh weight (FW) or root dry weight (DW) basis.

  • b

    Notes key: [A], uptake from ‘single salt’ solution containing both Zn2+ and Ca2+[B], uptake from a complete nutrient solution; [C] influx measurement ( 30 min); [D], uptake measurement (60–120 min); [E], calculated from Zn2+ depletion in uptake solution; [F], desorbed in cationic solution following uptake; [G], abrupt transitions between ‘phases’ described by single Michaelis–Menten functions; [H], a linear component was assumed to be cell wall binding.

  • c

    Vmax data recalculated assuming a root area/FW quotient of 20 cm2 g−1.

Schmid et al. (1965)0.5–1021.6714 FWNoNoNoNoNoExcised barley roots, [A], [C], [F], inhibition by 10 µm Cu
Chaudhry & Loneragan (1972a)0.5–103.1275 FWNoNoNoNoNoWheat seedlings, [A], [D], [F], noncompetitive inhibition by 250 µm Mg, Ba, Sr, Ca
Chaudhry & Loneragan (1972b)0.05–51.06.31 FWNoNoNoNoNoWheat seedlings, [A], [D], [F]
Chaudhry & Loneragan (1972c)0.5–103650.8 FWNoNoNoNoNoWheat seedlings, [A], pH 4, [D], [F], competitive inhibition by 1 µm Cu
Bowen (1973)8–500a) 184000 DWNoNoNoNoNoExcised sugarcane roots of (i) Zn-efficient [H53-263] and (ii) Zn-inefficient [H57-5174] clones, [A], [D], [F]
b) 10318 300 DW     
Brar & Sekhon (1976)0.5–25.34154 DWNoNoNoNoNoWheat plants, [A], [D], [F], competitive inhibition by 5 µm Cu
Hassan & van Hai (1976)0.01–500(a) 1.6 ± 1.0(a) 210 DW(a) 83 ± 24(a) 3880 DW(a) 120 ± 30(a) 7810 DWNo(a) Excised roots and (b) intact plants of sour orange, [B], [C], [G]
(b) 3.9 ± 3.2(b) 100 DW(b) 53 ± 18(b) 1790 DW(b) 150 ± 60(b) 7220 DW 
Veltrup (1978)1.5–13803.18 ± 1.8530 DW151 ± 43.810.6 DW490 ± 20018.2 DWNoBarley plants, [B], [D], rinse, [G], unaffected by 16 µm Cu
Bowen (1981)20–250165710 DW4220 400 DWNoNoNoExcised barley roots, [A], [C], [F], [G]
Homma & Hirata (1984)0.089–8.90.3728.7 FW5.4194.1 FWNoNoNoRice seedlings, [A], [D], [F]
Bowen (1986)10–500(a) 6(a) 2900 FWNoNoNoNoNoExcised root apices of (a) Zn-efficient [M101] and (b) Zn-inefficient [IR26] rice varieties, [A], [D], [F]
(b) 13(b) 5740 FW     
Mullins & Sommers (1986)0–101.5–2.22.9–4.0 FWcNoNoNoNoNoMaize plants, [B], [E]
Bowen (1987)10–500(a) 50(a) 7220NoNoNoNoNoExcised root apices of (a) Zn-efficient [Kewalo] and (b) Zn-inefficient [Sel 7625-2] tomato, [A], [C], [F], competitive inhibition by Cu
(b) 57(b) 2340     
Lasat et al. (1996)0.5–1008270 FWNoNoNoNo[H]Thlaspi caerulescens seedlings, [A], [C], [F]
Lasat et al. (1996)0.5–100660 FWNoNoNoNo[H]Thlaspi arvense seedlings, [A], [C], [F]
Rengel & Wheal (1997)0.03–3(a) 0.86 ± 0.09(a) 5.5 DWNoNoNoNoNoWheat seedlings of (a,b) Zn-efficient [Excalibur] and (c,d) Zn-inefficient [Gatcher] varieties, grown in (a,c) Zn-deficient or (b,d) Zn-replete solutions, [B], [E]
(b) 0.76 ± 0.06(b) 3.5 DW     
(c) 0.92 ± 0.14(c) 3.7 DW     
(d) 0.88 ± 0.07(d) 2.3 DW     
Rengel & Wheal (1997)0.03–3(a) 0.95 ± 0.10(a) 2.8 DWNoNoNoNoNoTriticum turgidum seedlings grown in (a) Zn-deficient and (b) Zn-replete solutions, [B], [E]
(b) 0.93 ± 0.09(b) 2.5 DW     
Wheal & Rengel (1997)0.0008–0.01230.002539 FWNoNoNoNoNoWheat plants, [B], [D], [E].
Hart et al. (1998)0–802.3 ± 0.4162 FWNoNoNoNo[H]Wheat seedlings, [A], [C], [F].
Hart et al. (1998)0–803.9 ± 0.5175 FWNoNoNoNo[H]Triticum turgidum, [A], [C], [F].
Santa-Maria & Cogliatti (1998)0.2–1022.3 ± 4.111 200 DWNoNoNoNoNoWheat plants, [B], [C], [F].
Hacisalihoglu et al. (2001)0.0001–80(a) 0.0006(a) 10.9 FW(a) 1.9  0.3(a) 446 FWNoNo[H]Wheat seedlings of (a,b) Zn-efficient [Dagdas] and (c,d) Zn-inefficient [BDME-10] varieties, grown in (a,c) Zn-deficient or (b,d) Zn-replete solutions, [A], [C], [F]
(b) 0.0012(b) 9.0 FW(b) 4.9 ± 1.7(b) 143 FW   
(c) 0.0023(c) 30.9 FW(c) 4.1 ± 1.5(c) 521 FW   
(d) 0.0007(d) 9.7 FW(d) 3.4 ± 1.1(d) 294 FW   
Hart et al. (2002)0.050–502.3 ± 0.3171 FWNoNoNoNo[H]Wheat seedlings [A], [C], [F], competitive inhibition by 10 µm Cd
Hart et al. (2002)0.050–503.3 ± 0.4166 FWNoNoNoNo[H]Triticum turgidum, [A], [C], [F], competitive inhibition by 10 µm Cd

Transport from epidermal and cortical cells to the root xylem can occur via the cytoplasmic continuum of cells, linked by plasmodesmata, from which Zn is pumped into the stelar apoplast (Lasat & Kochian, 2000). This ‘symplastic’ pathway is catalysed by plasma membrane and tonoplast transport activity. Zinc can also be delivered extracellularly to the stelar apoplast in regions where the Casparian band is not fully formed (White, 2001; White et al., 2002b). Apoplastic mineral fluxes are dominated by the cell wall cation exchange capacity (CEC), Casparian band formation and water flows (Sattelmacher, 2001). Both symplastic and apoplastic fluxes may contribute to net Zn fluxes to the shoot. If symplastic fluxes dominate, Zn accumulation in the shoot approximates the sum of unidirectional influx (φoc) of Zn to root cells, minus efflux (φco) and net vacuolar Zn sequesteration (i.e. unidirectional vacuolar influx, φcv, minus unidirectional vacuolar efflux, φvc) (Lasat & Kochian, 2000; White et al., 2002b). Thus, the maximal [Zn]shoot supported by symplastic root Zn fluxes can be simulated for different shoot : root FW ratios, relative growth rates (RGRs) and φoc values if one assumes steady-state conditions (White et al., 2002b). In Thlaspi caerulescens J. & C. Presl., which can accumulate > 30 000 µg Zn g−1 DW (see Section VI.3), the delivery of Zn to the root xylem exclusively via the symplast is kinetically challenging at high [Zn]ext (White et al., 2002b). Substantial symplastic Zn fluxes may occur at high [Zn]ext (White et al., 2002b). However, it is noteworthy that the Ca2+-transporting ATPase activity required to catalyze the export of Ca2+ alone from the symplast into the root stelar apoplast may exceed total membrane protein quotas in a typical dicot (White, 1998; White et al., 2002b). Thlaspi caerulescens has substantial shoot Ca, Mg and Zn concentrations (> 60,000 µg g−1 DW; Meerts et al., 2003; Dechamps et al., 2005). Thlaspi caerulescens also has altered Casparian band formation (Section VI.iii), and Cd2+ fluxes are substantially greater at the root tips in wheat, T. caerulescens and T. arvense L. (i.e. in regions where the Casparian band is not fully formed; Piñeros et al., 1998). Integrating Zn flux analyses with models to predict [Zn]ext may ultimately enable quantification of the relative symplastic and apoplastic root fluxes, provided fluxes and intracellular Zn distibutions of intact root cells can be determined accurately.

V. Zinc in plants

1. Zinc is an essential plant nutrient

The essentiality of Zn in plants was first shown in maize (Mazé, 1915), and subsequently in barley and dwarf sunflower (Sommer & Lipman, 1926). Early reports of severe Zn deficiency symptoms included impaired stem elongation in tomato (Skoog, 1940). Incipient Zn deficiency symptoms in tomato, remedied by resupply of Zn, included reduced protein and starch synthesis whilst sugar content was unaffected (Hoagland, 1948). Severe Zn deficiency symptoms have since been catalogued for many crops (Scaife & Turner, 1983; Marschner, 1995; Sharma, 2006; Fig. 1). Severe Zn deficiency is characterized by root apex necrosis (‘dieback’), whilst sublethal Zn deficiency induces spatially heterogeneous or interveinal chlorosis (‘mottle leaf’), the development of reddish-brown or bronze tints (‘bronzing’), and a range of auxin deficiency-like responses such as internode shortening (‘rosetting’), epinasty, inward curling of leaf lamina (‘goblet’ leaves) and reductions in leaf size (‘little leaf’). In most crops, the typical leaf Zn concentration ([Zn]leaf) required for adequate growth approximates 15–20 mg Zn kg−1 DW (Marschner, 1995).

Figure 1.

Zinc deficiency induced by prolonged flooding in rice. Typical symptoms include high plant mortality, stuntedness, leaf bronzing and a delay in flowering. (Photograph courtesy of Dr Matthias Wissuwa, Japan International Research Center for Agricultural Sciences, Tsukuba, Japan.)

Zinc is the most common crop micronutrient deficiency, particularly in high-pH soils with low [Zn]bss (Graham et al., 1992; White & Zasoski, 1999; Cakmak, 2002, 2004; Alloway, 2004). Notably, 50% of cultivated soils in India and Turkey, a third of cultivated soils in China, and most soils in Western Australia are classed as Zn-deficient. Zinc deficiency in crop production can be ameliorated through agronomy or genetic improvement. Early agronomic successes included the treatment of little leaf in peach orchards, using soil-applied FeSO4 fertilizers containing Zn impurities, and subsequently the treatment of mottle leaf in citrus orchards, rosetting in pecan and stunted pineapple growth using foliar sprays containing Zn (Hoagland, 1948). More recently, substantial arable crop responses to Zn fertilization have been reported in Australia, India, and Central Anatolia in Turkey, where wheat grain yields have increased by over 600% since the mid-1990s, with a concomitant annual economic benefit of US$100 million (Cakmak, 2004). It may also be possible to improve crop yields on Zn-deficient soils by exploiting genotypic differences in Zn uptake and tissue-use efficiency that exist within crop species (Rengel, 2001; Cakmak, 2002; Hacisalihoglu & Kochian, 2003; Alloway, 2004). For example, in rice, Zn uptake efficiency correlates with exudation rates of low-molecular-weight organic anions and a substantial proportion of the phenotypic variation in Zn uptake efficiency is under genetic control (Hoffland et al., 2006; Wissuwa et al., 2006).

2. Evolutionary aspects of shoot Zn concentration among angiosperms

Previously, a meta-analysis of 70 species from 35 studies in the literature implied that ancient evolutionary processes might impact on [Zn]shoot in angiosperms (Broadley et al., 2001). A much larger survey was undertaken here. Primary [Zn]shoot data, that is, Zn concentrations reported in leaf or nonwoody shoot tissues, were obtained from 1108 studies, contained in 204 published papers and three unpublished datasets (Tables S6–S9). These primary data define the largest set of interlinked studies, in which [Zn]shoot values are reported for more than two species in controlled experiments where each study contains more than one species common to another study. Thus, 365 species from 48 families and 12 key clades were sampled. Primary [Zn]shoot data were ln-transformed and a variance-components model was fitted to the data using residual maximum likelihood (REML) procedures (Broadley et al., 2001, 2003). Species, family and key clade variance components were estimated using a random model factor of (study + (key clade/family/species)) with no fixed model factor. Mean data for phylogenetic groups and significance tests (Wald tests) were conducted using (key clade/family/species) as a fixed factor, retaining (study + (key clade/family/species)) as a random factor. All analyses were performed using GenStat (Release 9.1.0.147; VSN International, Oxford, UK).

Study effects dominated the variance components because of the vast range of [Zn]ext across all studies and because of differences in the relationship between [Zn]ext and [Zn]shoot between species (Table S9). After removing study effects, key clade (Wald statistic = 592.15, d.f. = 10, P < 0.001) and family within key clade (Wald statistic = 298.71, d.f. = 36, P < 0.001), variance components accounted for 22.1% of the variation in [Zn]shoot among angiosperm species (Table S9). The analysis was repeated with Thlaspi and Arabidopsis genera excluded to remove the influence of Zn ‘hyperaccumulation’ (defined in Section VI.3) from the analysis. Adjusted key clade and family variance components accounted for 26.5% of the variation in [Zn]shoot (Table S9). After removing the effect of study, and with Thlaspi and Arabidopsis genera excluded, mean relative [Zn]shoot among key clades ranged from < 40 (Ericales) to > 100 (Caryophyllales and noncommelinoid monocotyledons; NB: units approximate µg Zn g−1 DW) (Table S9). Among well-replicated families, lower [Zn]shoot occurred in Linaceae (52, n = 29), Poaceae (59, n = 1527) and Solanaceae (66, n = 181), and higher [Zn]shoot occurred in Amaranthaceae (108, n = 214) and Salicaceae (195, n = 45) (Table S9). Compared with other essential elements, variation in [Zn]shoot manifesting above the family level is less than for Ca, K, Mg and Si (Broadley et al., 2004; Hodson et al., 2005), but greater than for N and P (Broadley et al., 2004). Furthermore, variation in [Zn]shoot manifesting above the family level is less than previously reported for [Zn]shoot among a limited dataset of 70 species (Broadley et al., 2001). However, despite the large study effects within this dataset, evidence of ancient evolutionary processes influencing [Zn]shoot can still be observed using this approach. A considerable additional sampling effort is required to explore this phenomenon further.

After removing study effects, > 70% of the remaining variation in [Zn]shoot occurs within-family. Thus, substantial differences in [Zn]shoot exist between and within genera and species. Several billion people worldwide have Zn-deficient diets, and this within-species genetic variation is providing a valuable genetic resource to select or breed crops with increased Zn concentrations in their edible portions, notably in several cereal, legume, root and leafy vegetable crops (Graham et al., 2001; Welch & Graham, 2004; Grusak & Cakmak, 2005; Vreugdenhil et al., 2005; White & Broadley, 2005; Ghandilyan et al., 2006). Accelerated breeding to increase the delivery of dietary Zn in crops may be possible following identification of intraspecific genetic variation in Zn composition. In common bean (Phaseolus vulgaris L.), seed Zn concentration ([Zn]seed) is a quantitative trait which can be mapped to genetic loci using quantitative trait loci (QTL) analyses (Ghandilyan et al., 2006). QTLs for [Zn]seed have also been mapped in A. thaliana (Vreugdenhil et al., 2004). Following the identification of QTL, candidate genes or loci can be resolved through fine mapping and map-based cloning, and this information could be used for gene-based selection or marker-assisted breeding strategies. An advantage of this strategy is that knowledge of the genes and/or chromosomal loci controlling [Zn]seed or [Zn]shoot in one plant species could be used in a different target crop species by exploiting gene homology and/or genome collinearity (Ghandilyan et al., 2006).

VI. Plant responses to elevated soil Zn

The response of plants to elevated [Zn]soil has generated a substantial literature, driven by the primary questions, ‘Can crops be grown safely and productively at elevated [Zn]soil?’, and, ‘How and why do certain taxa thrive at elevated [Zn]soil?

1. Zn toxicity in crops

Zinc toxicity in crops is far less widespread than Zn deficiency. However, Zn toxicity occurs in soils contaminated by mining and smelting activities, in agricultural soils treated with sewage sludge, and in urban and peri-urban soils enriched by anthropogenic inputs of Zn, especially in low-pH soils (Chaney, 1993). Toxicity symptoms usually become visible at [Zn]leaf > 300 mg Zn kg−1 leaf DW, although some crops show toxicity symptoms at [Zn]leaf < 100 mg Zn kg−1 DW (Chaney, 1993; Marschner, 1995), and toxicity thresholds can be highly variable even within the same species. For example, [Zn]leaf associated with a 50% yield reduction in radish ranged from 36 to 1013 mg kg−1 DW (Davies, 1993). Zn toxicity symptoms include reduced yields and stunted growth, Fe-deficiency-induced chlorosis through reductions in chlorophyll synthesis and chloroplast degradation, and interference with P (and Mg and Mn) uptake (Carroll & Loneragan, 1968; Boawn & Rasmussen, 1971; Foy et al., 1978; Chaney, 1993). Crops differ markedly in their susceptibility to Zn toxicity. In acid soils, graminaceous species are generally less sensitive to Zn toxicity than most dicots, although this is reversed in alkaline soils (Chaney, 1993). Among dicots, leafy vegetable crops are sensitive to Zn toxicity, especially spinach and beet, because of their inherent high Zn uptake capacity (Boawn & Rasmussen, 1971; Chaney, 1993). There is also genetic variation in sensitivity to Zn toxicity within species, including soybean (Glycine max L.; Earley, 1943; White et al., 1979a,b,c) and rice (Oryza sativa L.), in which QTLs impacting on sensitivity to Zn toxicity have recently been mapped (Dong et al., 2006).

2. Plant tolerance to elevated soil Zn (hypertolerance)

Numerous species of metallophyte thrive at a [Zn]soil that is toxic to most crop plants. Early studies on Zn hypertolerance focused on elevated [Zn]soil at mine sites and/or near corroded galvanized materials, such as electricity pylons (reviewed by Antonovics et al., 1971; Baker, 1987; Ernst et al., 1990, 1992; Macnair, 1993). Initially, Zn hypertolerance was thought to occur most frequently in species of Poaceae, Caryophyllaceae and Lamiaceae, although subsequent studies have shown no phylogenetic predisposition to the evolution of Zn hypertolerance (Baker, 1987). However, this hypothesis is difficult to test directly. At a species level, elegant theoretical and experimental frameworks have been developed to characterize the genetics of Zn hypertolerance (Macnair, 1990, 1993). Briefly, Zn hypertolerance tends to manifest as a dominant phenotype and spread rapidly in a population. However, dominance is not a fixed property of a gene but a measure of the phenotypic deviation of the heterozygote from the mean of the two homozygotes. Thus, G × E interactions will affect whether a gene confers a dominant or recessive phenotype under any particular condition, rendering the study of Zn tolerance nontrivial (Macnair, 1990, 1993). Nevertheless, Zn hypertolerance is likely to be under the control of a small number of major genes, as in Silene vulgaris (Moench) Garcke (Schat et al., 1996) and Arabidopsis halleri (L.) O’Kane & Al-Shehbaz (Macnair et al., 1999). In addition to dominance (i.e. interallelic/intragenic) effects, epistatic (i.e. intergenic) interactions have an impact on Zn hypertolerance, although these effects are more difficult to quantify (Macnair, 1993). Zinc-hypertolerant plants show fitness costs at low [Zn]soil, for example, suboptimal carbonic anyhdrase and nitrate reductase activity occurs in S. vulgaris (Ernst et al., 1992). In Silene dioica (L.) Clairv., pollen selection may accelerate reproductive isolation between adjacent populations which differ in Zn tolerance (Searcy & Mulcahy, 1985a,b,c).

At a whole-plant scale, natural Zn hypertolerance is thought to be conferred by Zn exclusion or by compartmentalization, for example, through mycorrhizal symbioses, altered root-to-shoot translocation or accumulation in older leaves (Baker, 1981; Ernst et al., 1992; Hall, 2002). Key subcellular processes enabling Zn hypertolerance are likely to be increased organic acid production and vacuolar compartmentalization, including increased Zn efflux across the plasma membrane (Ernst et al., 1992; Verkleij et al., 1998; Chardonnens et al., 1999; Clemens, 2001; Hall, 2002). Early hypotheses that cell-wall Zn binding, reduced membrane leakage or increased phytochelatin production increased Zn tolerance have not been supported by subsequent studies (Ernst et al., 1992; Harmens et al., 1993; Clemens, 2001; Schat et al., 2002).

In general, Zn hypertolerance does not segregate with other metal tolerance phenotypes, although Cd, Co, Cu, Cd, Ni and/or Pb cotolerance can occur (Gregory & Bradshaw, 1965; Wu & Antonovics, 1975; Cox & Hutchinson, 1980; Symeonidis et al., 1985; Macnair, 1990, 1993; Brown & Brinkmann, 1992; Schat & Vooijs, 1997). Cotolerance can be either pleiotropic (i.e. strict cotolerance) or the result of multiple hypertolerance mechanisms. Multiple hypertolerances arise through nonrandom association of favourable alleles at two or more loci (i.e. linkage disequilibrium), potentially in response to the co-occurrence of several metals in soils, gene flow from adjacent sites or seed transport between different metalliferous sites (Macnair, 1993; Schat & Vooijs, 1997). For example, Zn/Co/Ni cotolerance phenotypes cosegregated in Silene vulgaris, implying a pleiotropic effect, whilst Zn/Cd/Cu cotolerance phenotypes segregated independently, implying multiple mechanisms (Schat & Vooijs, 1997). Zinc/Cu cotolerance phenotypes also segregated independently in populations of Agrostis stolonifera L. (Wu & Antonovics, 1975) and Mimulus guttatus DC. (Macnair, 1990). Zinc-hypertolerant cultivars of grass species have been successfully bred and used to revegetate soils contaminated with Zn, Pb and other heavy metals, following mining activities, including Festuca rubra L. cv. Merlin and Agrostis capillaris L. cv. Groginan for calcareous and acid wastes, respectively (Smith & Bradshaw, 1979; Whiting et al., 2005).

3. Zinc hyperaccumulation

The ability of some Zn-hypertolerant metallophytes to accumulate exceptional concentrations of Zn in their aerial parts was probably first reported among ‘galmei’ (calamine) flora of the Aachen region on the border of Belgium and Germany, where the presence of > 1% Zn in the plant ash of Viola calaminaria (Ging.) Lej. was recorded in 1855 (Reeves & Baker, 2000). A [Zn]shoot > 1% DW in Thlaspi alpestre L. (= T. caerulescens) was reported shortly thereafter in 1865 (Reeves & Baker, 2000). Zinc hyperaccumulation has since been defined as the occurance of > 10 000 µg Zn g−1 DW in the aerial parts of a plant species when growing in its natural environment (Baker & Brooks, 1989). These species (Table 3) have come to dominate the literature, in part because of the desire to transfer the character into crop species for use in phytoremediation, phytomining and biofortification (Chaney, 1983; Baker & Brooks, 1989; Brooks, 1998; Salt et al., 1998; Guerinot & Salt, 2001; Macnair, 2003; Krämer, 2005a). Typically, 10–20 species are reported to be Zn hyperaccumulators, with a smaller number of these able to accumulate Cd to very high concentrations as well. This raises the intriguing question: how often has the Zn hyperaccumulation character evolved? The answer is uncertain because: (i) 3000 µg Zn g−1 DW might be a more suitable evolutionary definition of extreme [Zn]shoot based on the frequency distribution of values observed within the genus Thlaspi s.l., which contains most Zn hyperaccumulators (Reeves & Brooks, 1983; Reeves & Baker, 2000); (ii) records for several nonbrassicaceous Zn hyperaccumulators are uncertain (Macnair, 2003; R. D. Reeves, pers. comm.) (NB: plant samples collected from metal-rich substrates can easily become contaminated by soil particles); and (iii) taxonomic uncertainties and the use of synonyms can affect the identification of field and herbarium samples.

Table 3.  Plant species whose [Zn]shoot has been observed to exceeds 0.3% DW (unless stated) when grown under natural conditions
SpeciesaPotential synonymsaFamily (Order)LocalityMaximum [Zn]shoot (% DW) observed).References and Comments
Acer pseudoplatanus L.Sapindaceae (Sapindales)UK0.35Johnston & Proctor (1977)
Arenaria patula Michx.Minuartia patula (Michx.) Mattf.Caryophyllaceae (Caryophyllales)USA1.31Brooks (1998). Uncertain record (Macnair, 2003; R. D. Reeves, pers. comm.)
Arabidopsis arenosa (L.) LawalréeCardaminopsis arenosa (L.) HayekBrassicaceae (Brassicales)France0.52Reeves et al. (2001)
Arabidopsis halleri (L.) O’Kane & Al-ShehbazArabis gemmifera Makino, Cardaminopsis halleri (L.) Hayek, C. ovirensis (Wulf.) O. SchwarzBrassicaceaeFrance2.07R. D. Reeves, pers. comm.
Arabidopsis thaliana (L.) Heynh.Arabis thaliana L., Sisymbrium thalianum (L.) J. Gay & Monnard, Stenophragma thalianum (L.) Čelak.BrassicaceaeUSA2.67Reeves (1988)
Biscutella laevigata L.Biscutella alsatica Jord., B. austriaca Jord., B. longifolia Vill., B. lucida Balb. ex DC., B. sempervirens L., B. varia Dumort., B. variegata Boiss. & Reut., B. vincentina (Samp.) Rothm. ex GuineaBrassicaceaeFrance0.41R. D. Reeves, pers. comm.
Cochlearia pyrenaica DC.Cochlearia officinalis L. subsp. pyrenaica (DC.) Rouy & FoucaudBrassicaceaeUK0.53Reeves (1988)
Dianthus sp.CaryophyllaceaeFrance0.49R. D. Reeves, pers. comm.
Dichapetalum gelonioides (Roxb.) Engl.Chailletia gelonioides (Roxb.) J. D. Hook., Dichapetalum howii Merrill & Chun., Moacurra gelonioides Roxb.Dichapetalaceae (Malpighiales)Indonesia, Malaysia, Philippines3.00Reeves & Baker (2000)
Galium mollugo L.Galium album Mill., G. cinereum All., G. corrudifolium Vill., G. elatum Thuill., G. insubricum Gaudin, G. kerneranum Klokov, G. lucidum All., G. neglectum Le Gall ex Gren., G. tyrolense Willd.Rubiaceae (Gentianales)France0.30Reeves et al. (2001)
Gomphrena canescens R. Br.Amaranthaceae (Caryophyllales)Australia0.90Nicolls et al. (1965)
Haumaniastrum katangense (S. Moore) Duvign. & Plancke.Lamiaceae (Lamiales)D. R. of the Congo1.98Brooks (1998). Uncertain record (Paton & Brooks, 1996; Macnair, 2003)
Minuartia verna (L.) HiernAlsine verna (L.) Wahlenb., Arenaria verna L., Minuartia caespitosa (Ehrh.) DegenCaryophyllaceaeYugoslavia1.14Various studies cited in Reeves & Baker (2000)
Noccaea boeotica F. K. Mey.BrassicaceaeGreece0.31R. D. Reeves, pers. comm.
Noccaea eburneosa F. K. Mey.Noccaea salisii (Brugger) F. K. Mey, Thlaspi salisii BruggerBrassicaceaeSwitzerland1.05Reeves & Brooks (1983), taxonomic status uncertain (R. D. Reeves, pers. comm.)
Polycarpaea synandra F. Muell.CaryophyllaceaeAustralia0.70Reeves & Baker (2000)
Rumex acetosa L.Polygonaceae (Caryophyllales)UK1.10Johnston & Proctor (1977)
Rumex acetosella L.Acetosella vulgaris Fourr., Rumex acetoselloides Balansa, R. angiocarpus Murb., R. fascilobus Klokov, R. multifidus L., R. pyrenaicus Pourr. ex Lapeyr., R. salicifolius auct., R. tenuifolius (Wallr.) Á. Löve PolygonaceaeFrance0.31Reeves et al. (2001)
Sedum alfredii HanceCrassulaceae (Saxifragales)China0.50Yang et al. (2002)
Silene vulgaris (Moench) GarckeSilene angustifolia Mill., S. campanulata Saut., S. cucubalus Wibel, S. inflata Sm., S. latifolia (Mill.) Britten & Rendle, non Poir., S. tenoreana Colla, S. venosa Asch., S. vulgaris (Moench) GarckCaryophyllaceaeUSA0.47Brooks (1998)
Thlaspi apterum Velen.Noccaea aptera (Velen.) F. K. Mey.BrassicaceaeBulgaria0.31Reeves & Brooks (1983). Uncertain record (R. D. Reeves, pers. comm.)
Thlaspi alpinum CrantzNoccaea alpestris (Jacq.) Kerguélen, N. sylvia (Gaudin) F. K. Mey., T. sylvium GaudinBrassicaceaeFrance0.54Reeves et al. (2001)
Thlaspi brachypetalum Jord.Noccaea brachypetala (Jord.) F. K. Mey.BrassicaceaeFrance1.53Reeves & Brooks (1983)
Thlaspi brevistylum (DC.) Mutel.Noccaea brevistyla Steud.BrassicaceaeCorsica0.31Taylor (2004)
Thlaspi bulbosum Spruner ex Boiss.Raparia bulbosa (Boiss.) F. K. Mey.BrassicaceaeGreece1.05Brooks (1998)
Thlaspi caerulescens J. & C. Presl.Noccaea arenaria (J. E. Duby) F. K. Mey., N. caerulescens (J. & C. Presl) F. K. Mey., N. occitanica (Jord.) F. K. Mey., Thlaspi alpestre L., T. arenarium Jord., T. caerulescens subsp. caerulescens, T. caerulescens subsp. calaminare (Lej.) Lej. & Court, T. caerulescens subsp. occitanicum (Jord.) M. Laínz, T. caerulescens subsp. tatrense (Zapał.) Dvořáková, T. gaudinianum Jord., T. huteri A. Kern., T. mureti Gremli, T. occitanicum Jord., T. pratulorum Gand., T. rhaeticum Jord., T. salisii Brugger, T. suecicum Jord., T. sylvestre Jord., T. tatrense Zapał., T. virgatum Gren. & Godr., T. villarsianum Jord., T. vogesiacum Jord., T. vulcanorum LamotteBrassicaceaeW. and C. Europe4.37Various studies (Reeves & Baker, 2000). Also a Cd hyperaccumulator
Thlaspi cepaeifolium (Wulfen) W. D. J. KochNoccaea cepaeifolia (Wulfen) Rchb., N. limosellifolia (Burnat) F. K. Mey., N. rotundifolia (L.) Moench, Thlaspi cepaeifolium subsp. rotundifolium (L.) Greuter & Burdet, T. limosellifolium Reut. ex Rouy & Fouc., T. rotundifolium (L.) Gaudin subsp. cepaeifolium (Wulfen) Rouy & Fouc.BrassicaceaeItaly2.10Various studies (Reeves & Baker, 2000)
Thlaspi epirotum Hal.Noccaea epirota (Hal.) F. K. Mey.BrassicaceaeGreece<0.30Reeves & Brooks (1983)
Thlaspi goesingense Hal.Noccaea goesingensis (Hal.) F. K. Mey., Thlaspi tymphaeum Hausskn., T. umbrosum Waisb.BrassicaceaeAustria0.38Reeves & Baker (1984)
Thlaspi graecum Jord.Noccaea graeca (Jord.) F. K. Mey., Thlaspi taygeteum Boiss.BrassicaceaeGreece<0.30Reeves & Brooks (1983)
Thlaspi kovatsii Heuff.Noccaea kovatsii (Heuffel) F. K. Mey., Thlaspi affine Schott & Kotschy, T. avalanum Panč., T. jankae Kern., T. trojagense Zapał.BrassicaceaeBulgaria0.49Reeves & Brooks (1983)
Thlaspi magellanicum Pers.Noccaea magellanica (Pers.) J. HolubBrassicaceaeArgentina0.39Reeves (1988)
Thlaspi montanum L.Noccaea alpestris (Jacq.) Kerguélen subsp. sylvium (Gaudin), N. montana (L.) F. K. Mey., Thlaspi lotharingum Jord.BrassicaceaeUSA0.43Hobbs & Streit (1986)
Thlaspi ochroleucum Boiss. & Heldr.Noccaea lutescens (Velen.) F. K. Mey., N. ochroleuca (Boiss. & Heldr.) F. K. Mey., N. phrygia (Bornm.) F. K. Mey., N. rhodopensis F. K. Mey., N. versicolor (Stoj. & Kitanov) F. K. Mey., Thlaspi balcanicum Janka, T. heterochroum Boiss., T. lutescens Velen., T. phrygium Bornm.BrassicaceaeTurkey0.63Reeves & Brooks (1983), Reeves (1988)
Thlaspi parviflorum A. Nels.Noccaea parviflora (A. Nels.) HolubBrassicaceaeUSA0.31Reeves et al. (1983)
Thlaspi pindicum Hausskn.Noccaea pindica (Hausskn.) J. Holub, N. tymphaea (Hausskn.) F. K. Mey., Thlaspi tymphaeum Hausskn.BrassicaceaeGreece<0.10Taylor & Macnair, 2006. Note, plants collected from serpentine soils with low [Zn]soil. [Zn]shoot >1.00% DW observed under laboratory conditions.
Thlaspi praecox Wulf.Noccaea praecox (Wulf.) F. K. Mey, Thlaspi affine Schott & Kotschy ex Bioss.BrassicaceaeBulgaria2.10Brooks (1998). Cd hyperaccumulator (Vogel-Mikušet al., 2005)
Thlaspi stenopterum Boiss. & Reut.Noccaea stenoptera (Boiss. & Reut.) F. K. Mey.BrassicaceaeSpain1.60Brooks (1998)
Thlaspi viridisepalum (Podp.) Greuter & BurdetNoccaea viridisepala (Podp.) F. K. Mey.BrassicaceaeBulgaria0.63Reeves & Brooks (1983)
Viola calaminaria (Gingins) Lej.Viola tricolor L.Violaceae (Malpighiales)Germany1.00Brooks (1998). Note, uncertain record (Macnair, 2003; R. D. Reeves, pers. comm.)

The genus Thlaspi L. s.l. is likely to be polyphyletic and its taxonomy is controversial. Seed-coat anatomical and sequence data have thus been used to split the genus into several alternative genera, including Thlaspi s.s., Vania and a clade containing Thlaspiceras, Noccaea, Raparia, Microthlaspi and Neurotropis (Meyer, 1973, 1979; Mummenhoff & Koch, 1994; Mummenhoff et al., 1997; Koch & Mummenhoff, 2001; Koch & Al-Shehbaz, 2004). High [Zn]shoot is probably a general feature of Noccaea and its sister clade Raparia (Macnair, 2003; Taylor, 2004), but not of Thlaspiceras, which nevertheless contains Zn-hypertolerant species (e.g. Thlaspiceras oxyceras (Boiss.) F. K. Mey; Peer et al., 2003), nor of the more distantly related nonZn-hypertolerant Microthlaspi and Neurotropis clades. Thus, the high [Zn]shoot character most likely evolved at the base of the Noccaea/Raparia clade, or less likely at the base of the Noccaea/Raparia/Thlaspiceras clade with a subsequent reversion to the low [Zn]shoot character in Thlaspiceras (Macnair, 2003; Taylor, 2004). Intriguingly, since Ni hyperaccumulation also evolved at the base of the Noccaea/Raparia/Thlaspiceras clade, high [Zn]shoot may be a modification of the Ni hyperaccumulation character (Taylor, 2004). Nickel hyperaccumulation is more common than Zn hyperaccumulation among angiosperms (> 300 species, in c. 34 families), although > 80% of temperate Ni hyperaccumulators are Brassicaceae species (Reeves & Baker, 2000; Borhidi, 2001). The only other Brassicaceae species with an unequivocally high [Zn]shoot character is Arabidopsis halleri. Thus, Zn hyperaccumulation has probably only arisen during two relatively recent evolutionary events within the Brassicaceae, and possibly on very few isolated occasions elsewhere in the angiosperms. The following sections focus primarily on Thlaspi caerulescens.

Genetics of Zn hyperaccumulation Thlaspi caerulescens is a short-lived, self-compatible biennial/perennial species which hyperaccumulates Zn/Cd/Ni. It occurs on calamine, serpentine (derived from ultramafic Mg and Fe-rich rocks, also enriched with Co, Cr and Ni), and nonmineral soils, with a wide distribution in central, northern and western Europe (Reeves & Brooks, 1983; Baker & Brooks, 1989; Reeves et al., 2001). It is functionally nonmycorrhizal (Regvar et al., 2003). In T. caerulescens, Zn hyperaccumulation and hypertolerance are constitutive species-level traits although inter- and intrapopulation variations in [Zn]shoot and/or Zn tolerance occur (Mathys, 1977; Ingrouille & Smirnoff, 1986; Baker et al., 1994; Lloyd-Thomas, 1995; Pollard & Baker, 1996; Meerts & Van Isacker, 1997; Escarréet al., 2000; Reeves et al., 2001; Pollard et al., 2002; Assunção et al., 2003a,b, 2006; Frérot et al., 2003, 2005; Macnair, 2003; Roosens et al., 2003; Zha et al., 2004; Molitor et al., 2005). Populations from soils with high-[Zn]soil are frequently described as ‘metallicolous’ with an explicit adaptive connotation. Notably, outcrossing rates of c. 5.25% have been reported (Riley, 1956), although rates are population-specific and allozyme variation and pollen : ovule ratios indicate that low-[Zn]soil populations are more self-fertile than high-[Zn]soil populations (Koch et al., 1998; Dubois et al., 2003). Conversley, in nonhyperaccumulator species, metal-hypertolerant populations are more self-fertile than nontolerant populations (Antonovics, 1968, 1972; Ducousso et al., 1990).

When T. caerulescens are grown under identical experimental conditions, Zn tolerance is generally greater, and [Zn]shoot is lower in high-[Zn]soil populations than in low-[Zn]soil populations. For example, [Zn]shoot was higher in field-sampled high-[Zn]soil populations than in low-[Zn]soil populations from the UK, but half-sib families of high-[Zn]soil populations had lower [Zn]shoot than low-[Zn]soil populations when grown subsequently in hydroponics (Pollard & Baker, 1996). Among a wider sample of European populations, [Zn]shoot, but not [Zn]xylem (ranges 63–93 and 610–700 µm at 10 and 100 µm[Zn]ext, respectively), varied similarly (Roosens et al., 2003). In soil-based experimental studies lasting up to 1 yr, Zn tolerance was lower and [Zn]shoot higher in low-[Zn]soil populations than in high-[Zn]soil populations (Meerts & Van Isacker, 1997; Escarréet al., 2000). Variation in [Zn]shoot between and within half-sib families was significant, but less than variation between populations, consistent with the self-fertilizing character of T. caerulescens. Notably, genetic variation in [Zn]shoot can occur within a few kilometres (Molitor et al., 2005). In A. halleri, Zn hyperaccumulation and hypertolerance are also constitutive traits at the species level, although interspecific crosses between A. halleri and A. lyrata ssp. petraea (L.) O’Kane & Al-Shehbaz reveal that Zn hyperaccumulation and hypertolerance are genetically independent (Macnair et al., 1999). Again, low-[Zn]soil populations accumulate more Zn than high-[Zn]soil populations of A. halleri under identical experimental conditions (Bert et al., 2000, 2002).

There is a substantial G × E interaction impacting on [Zn]shoot. For example, Assunção et al. (2003a,b, 2006) studied low-[Zn]soil populations of T. caerulescens from a nonmetalliferous (Lellingen, Luxembourg (LE)) site and a serpentine (Monte Prinzera, Italy (MP)) site, and high-[Zn]soil populations from two calamine sites (La Calamine, Belgium (LC) and Ganges, France (GA)). In field samples, [Zn]shoot decreased GA ≥ LC > LE > MP. In hydroponics, [Zn]shoot decreased MP > GA > LE > LC and Zn tolerance decreased GA = LC > MP > LE. A controlled intraspecifc cross was made from LE and LC; Zn accumulation segregated in F3s with a continuous phenotypic distribution indicative of a polygenic trait. However, the genetic independence of Zn tolerance and accumulation in T. caerulescens could not be confirmed because of the negative correlation between the two traits. A genetic linkage map was subsequently constructed for this population and two QTLs were mapped for [Zn]root (on chromosomes 3 and 5), although no QTLs were identified for [Zn]shoot (Assunção et al., 2006). However, in an F2 population obtained from an LC × GA cross, three QTLs explained 44.5% of the phenotypic variance in [Zn]shoot, with positive effects on [Zn]shoot arising as a result of both LC and GA alleles (Deniau et al., 2006). Similarly, in controlled crosses from high- and low-[Zn]soil populations of T. caerulescens (Frérot et al., 2003, 2005), F1 progeny with at least one parent from a high-[Zn]soil population were more sensitive to Zn deficiency, but F1 and F2 progenies were more tolerant to high [Zn]ext and had lower [Zn]shoot than progenies from exclusively low-[Zn]soil population crosses. Zha et al. (2004) also studied [Zn]shoot in F2 progenies of a controlled cross between a high (Ganges, southern France) and a low (Prayon, Belgium) Cd accumulator. When plants were grown in a Zn-supplemented compost, Ganges accumulated c. 50% higher [Zn]shoot than Prayon and parental frequency distributions overlapped. In the F2s, [Zn]shoot had a continuous distribution. There was significant transgression below the distribution of Prayon parents, but not above Ganges parents. Conversely, in hydroponics in the presence of higher [Cd]ext, [Zn]shoot was lower in Ganges than in Prayon, perhaps because of a toxic effect of Cd on the growth of Prayon. Again, the F2s showed a continuous distribution for [Zn]shoot and significant transgression below the lower limit of Ganges parental distributions, but not above Prayon (Zha et al., 2004). Intriguingly, other studies have also shown that high [Cd]ext inhibits Zn accumulation in Ganges, but not in Prayon, whilst high [Zn]ext inhibits Cd accumulation in Prayon, but not in Ganges populations (Lombi et al., 2001; Zhao et al., 2002; Roosens et al., 2003). In two species endemic to serpentine soils (Thlaspi pindicum Hausskn. and T. alpinum Crantz var. sylvium Gaudin), [Zn]shoot was higher when Ni was also present in the hydroponic solution (Taylor & Macnair, 2006).

Compartmentalization of Zn in hyperaccumulators  Generally, in Zn hyperaccumulators, [Zn]shoot > [Zn]root by up to 10-fold, although this depends on [Zn]ext (Shen et al., 1997; Zhao et al., 2000; Roosens et al., 2003). Vázquez et al. (1992, 1994) used scanning transmission (STEM) electron microscopy/energy-dispersive X-ray microanalysis (EDXMA) on freeze-substituted tissues to study cellular Zn distribution in T. caerulescens. Vacuoles of leaf epidermal and mesophyll cells, and leaf cell walls contained 296, 80 and c. 110 µg Zn g−1 DW, respectively, in plants grown at low [Zn]ext. At high [Zn]ext, vacuoles of leaf epidermal cells and their cell walls contained up to 13 600 and 929 µg Zn g−1 DW, respectively. Vacuoles of leaf mesophyll cells and their cell walls contained up to 4610 and 333 µg Zn g−1 DW, respectively. Zinc also accumulated in leaf subepidermal cells. In root epidermal and subepidermal cells, the Zn concentration ratio was approx. 1 (56 µg Zn g−1 DW) : 1.7 : 3 : 3.5 (vacuole centre : vacuole periphery : cell wall : intercellular space) at low [Zn]ext. At high [Zn]ext, background vacuolar Zn concentration was 175 µg Zn g−1 DW; scattered, electron-dense deposits of up to 18 300 µg Zn g−1 DW (likely to be artefacts of sample preparation; Küpper et al., 1999; Frey et al., 2000; Ma et al., 2005) were also reported. The Zn concentration ratio was c. 1 : 0.5 : 1 (background vacuole : cell wall : intercellular space). Thus, root cell wall Zn concentrations were, in general, similar at different values of [Zn]ext but vacuolar Zn concentrations differed. Küpper et al. (1999) used direct microcapillary sampling and scanning electron microscopy (SEM)-EDXMA on freeze-fractured T. caerulescens leaves. Epidermal and mesophyll cell saps contained 385 and 60 mm, respectively, but the latter represented c. 60% of the leaf Zn content. Zinc did not accumulate in subepidermal cells. Vacuolation promoted Zn accumulation, that is, Zn concentrations were lower in younger leaves and correlated with epidermal cell length (Küpper et al., 1999). Arabidopsis halleri has thinner leaves and smaller epidermal cells (length 8–30 µm) than T. caerulescens (10–100 µm) and also has trichomes (Küpper et al., 2000; Zhao et al., 2000). In contrast to T. caerulescens, SEM-EDXMA on freeze-fractured A. halleri tissues revealed Zn to be more uniformly distributed across the leaf, although trichome bases were substantially enriched with Zn (up to 1 m; Küpper et al., 2000; Zhao et al., 2000). Mesophyll cells had two- to threefold higher Zn concentrations than smaller-vacuoled epidermal cells and, at high [Zn]ext, relative Zn enrichment of the mesophyll cells was greater than in the trichomes. Consistent with EDXMA studies, not all leaf protoplasts of A. halleri accumulate Zn, although all leaf protoplasts of A. halleri (and T. caerulescens) are Zn-hypertolerant (Marquès et al., 2004). In A. halleri roots, Zn-phosphate precipitation in the rhizodermal outer wall prevented substantial Zn accumulation in other root compartments (Küpper et al., 2000).

Frey et al. (2000) quantified subcellular Zn distribution in ultrathin (100 nm) cryosections of T. caerulescens using STEM-EDXMA. Zinc localized to upper and lower leaf epidermal cells. Leaf epidermal cells contained 74 305, 11 577 and 3205 µg Zn g−1 DW in their vacuoles, cell walls/apoplast and cytoplasm, respectively. In contrast, leaf mesophyll (327, 9353, 262), guard (1439, 8438, 589) and subsidiary (3009, 9615, 2420) cells and root cortical cells (262, 589, 262) contained substantially less Zn (values given in parentheses represent vacuole, cell wall/apoplast and cytoplasm compartments, respectively, in µg Zn g−1 DW). Thus, a substantial fraction of total shoot Zn content was localized to the cell wall/apoplast, for example, 79% of the mesophyll cell Zn quota. Studies using the Zn-specific dye Newport green diacetate confirm that extremely low cytoplasmic Zn concentrations are maintained in Zn hyperaccumulators (Marquès et al., 2004). In direct analyses of mesophyll and epidermal protoplasts, mesophyll vacuoles and peeled leaf fractions, epidermal : mesophyll Zn ratios were reportedly approx. 2.5 : 1, with the mesophyll accounting for 65% of the leaf Zn quota (Ma et al., 2005). However, only 23% of the leaf mesophyll cell Zn content was localized to the cell wall/apoplast. Similarly, c. 9–35% of mesophyll cell Cd quota has been attributed to apoplastic compartments (Cosio et al., 2005; Ma et al., 2005). Although different quantification methods will lead to discrepancies, compartmentalization of Zn in leaf cell vacuoles and cell walls is clearly an important facet of Zn hyperaccumulation.

In what form is Zn stored in hyperaccumulators?  In T. caerulescens and A. halleri, up to 80% of shoot Zn is soluble in water or weak acids (Tolràet al., 1996; Zhao et al., 1998, 2000; Ma et al., 2005). In contrast to many crop species, insoluble Zn : P complexes such as Zn3(PO4)2 and Zn-phytates are not present in significant quantities (Zhao et al., 1998, 2000; Sarret et al., 2002). In roots, insoluble fractions make up a much larger proportion of the Zn content. For example, 25–57% of root Zn content is soluble but this declines at high [Zn]ext, whilst insoluble Zn correlates with insoluble P in the stoichiometric ratio expected of Zn3(PO4)2, that is, 0.27 P : Zn (Zhao et al., 1998, 2000). In T. caerulescens leaf cells, millimole equivalents (meq) of the major cations (Ca + K + Mg + Zn) ranged from 167 to 562 meq except in the vacuoles of mesophyll (756 meq) and epidermal (1100 meq) cells (Frey et al., 2000) and soluble vacuolar Zn concentrations did not associate with P, S or Cl (Küpper et al., 1999; Frey et al., 2000). Phytochelatins, small cystein-rich peptides, do not have an important role in binding Zn in hyperaccumulators, or in conferring Zn hypertolerance in general (Ernst et al., 1992; Harmens et al., 1993; Küpper et al., 2000, 2004; Clemens, 2001; Schat et al., 2002; Callahan et al., 2006; Wójcik et al., 2006). Further, the role of many other organic/amino acids, peptides and proteins (e.g. metallothioneins and the phytosiderophore precursor nicotianamine), which can also bind Zn, in Zn hyperaccumulation is not yet known (Callahan et al., 2006). However, since inorganic cation and organic anion equivalents correlate significantly in T. caerulescens shoots (Tolràet al., 1996), and since carboxylic acids, primarily malate, citrate and oxalate, and amino acids are abundant in plant materials, their role in Zn hyperaccumulation has been studied widely (Tolràet al., 1996; Shen et al., 1997; Zhao et al., 1998, 2000; Salt et al., 1999; Küpper et al., 2000, 2004; Sarret et al., 2002; Ma et al., 2005; Callahan et al., 2006; Wójcik et al., 2006).

In T. caerulescens shoots, concentrations of malate (164–248 µmol g−1 FW) > citrate (50–87 µmol g−1 FW) > succinate (23–38 µmol g−1 FW) > oxalate (1.9–4.9 µmol g−1 FW) > fumarate (0.31–1.25 µmol g−1 FW) > cis-aconitate (0.17–0.24 µmol g−1 FW), trans-aconitate (< 0.06 µmol g−1 FW); formate and acetate are detectable only at high [Zn]ext (Tolràet al., 1996). Soluble Zn, malate and oxalate are correlated in T. caerulescens shoots, although organic acid concentrations are constitutively high (Tolràet al., 1996; Shen et al., 1997; Wójcik et al., 2006), as they are in A. halleri (Zhao et al., 2000). The molar ratios of malate : Zn in T. caerulescens (4.8–72, (Tolràet al., 1996), 9.7–23.6 (Wójcik et al., 2006)) are sufficient to support a Zn-malate shuttle hypothesis (reviewed by Ernst et al., 1992), that is, the transport of Zn-malate across the tonoplast and dissociation and subsequent Zn2+ binding to stronger chelators such as citrate or oxalate, and export of malate back across the tonoplast to the cytosol. However, malate does not have a strong affinity for Zn (Tolràet al., 1996). Whilst oxalate : Zn molar ratios are low in T. caserulescens (0.1–0.68; Tolràet al., 1996), molar ratios of citrate : Zn are 2.1–5.9 in T. caerulescens (Wójcik et al., 2006) and 0.37–49.0 in A. halleri (Zhao et al., 2000). In T. caerulescens roots, organic acid concentrations are substantially lower than in shoots; malate (2.1–10 µmol g−1 FW) = citrate (3.2–16.5 µmol g−1 FW) = succinate (1.8–10.6 µmol g−1 FW) > fumarate (0.027–0.18 µmol g−1 FW) > cis-aconitate (< 0.07 µmol g−1 FW) = trans-aconitate (< 0.07 µmol g−1 FW). Acetate and oxalate (1.4 µmol g−1 FW) were only detectable at high [Zn]ext (Tolràet al., 1996). Whilst some studies report little correlation between Zn accumulation and altered root organic acid status (Tolràet al., 1996), Zn might stimulate citrate production in T. caerulescens roots (Shen et al., 1997).

Zinc coordination to O, N, S and His ligands can be predicted using noninvasive X-ray absorption spectroscopy (XAS) and extended X-ray absorption fine structure (EXAFS) analysis (Salt et al., 1999; Sarret et al., 2002; Küpper et al., 2004; Callahan et al., 2006). In T. caerulescens, up to 70% of root Zn may be associated with His, the remaining 30% with the cell wall, although it is difficult to discriminate between Zn(His)2 and Zn2+ bound to O ligands, potentially representing hydroxyl groups of water, using these techniques (Salt et al., 1999; Callahan et al., 2006). In xylem saps, 21% of Zn was found to be associated with Zn citrate, the reminder probably being free Zn2+. In shoots, 38, 9, 16 and 12% of total Zn was associated with citrate, oxalate, His and the cell wall, respectively, with 26% as free Zn2+ (Salt et al., 1999). No evidence of Zn-malate coordination was found. In other studies, Zn was predominantly coordinated with O ligands, potentially representing hydroxyl groups of water, or potentially carboxyl groups of malate, citrate or other organic acids (Küpper et al., 2004). Stronger Zn binding to His occurred in young leaves, potentially to avoid toxicity, and in senescing leaf tissues, which could be as a result of the breakdown in cell compartmentation. No associations between Zn and S ligands such as phytochelatins, metallothioneins or other Cys-rich peptides were observed (Küpper et al., 2004). In A. halleri shoots, Zn was primarily (octahedrally) coordinated with malate. In the trichomes, a secondary unidentified organic Zn compound was present, tetrahedrally coordinated and complexed to carboxyl and/or hydroxyl functional groups (Sarret et al., 2002). In A. lyrata ssp. petraea, shoot Zn was coordinated primarily with phosphate.

Two unusual root properties of Zn hyperaccumulators  Two remarkable features of T. caerulescens roots may be linked to its ability to hyperaccumulate Zn. First, a zincophilic root foraging response to heterogeneous [Zn]soil has been shown, analogous to the exploitation of spatially heterogeneous soil macronutrients (Schwartz et al., 1999; Whiting et al., 2000; Haines, 2002). Increases in root biomass (including root length and root hair production) occur in high-Zn-containing patches compared with adjacent Zn-deficient patches. These responses are not thought to be constitutive at the species level (Whiting et al., 2000; Haines, 2002) and warrant further genetic and molecular investigations. Second, a ‘peri-endodermal’ layer of cells with irregularly thickened inner tangential walls extending to < 1 mm from the root tip has been reported recently in T. caerulescens (Zelko et al., in press; Figs 2, 3). This layer is composed of secondary cell walls impregnated by suberin/lignin, forming a compact cylinder surrounding the endodermis from the outer side. This layer is not seen in T. arvense (Zelko et al., in press) or A. thaliana (van de Mortel et al., 2006) when compared with T. caerulescens. The development of the endodermis also differs between T. caerulescens and T. arvense (Zelko et al., in press). In T. caerulescens, Casparian bands (the first stage of endodermal development) form < 1 mm from the root tip, and suberin lamellae (the second stage of endodermal development) are formed in all endodermal cells c. 5–6 mm from the apex. In T. arvense, Casparian bands develop c. 2 mm from the root tip and suberin lamellae are formed in all endodermal cells > 10 mm from the root tip. Although a similar feature to this peri-endodermal layer was observed by early anatomists in some Brassicaceae, being described as ‘réseau sus-endodermique’ in 1887 (van Tieghem, 1887; Zelko et al., in press), the precise function of this layer of cells and its impact on apoplastic and symplastic fluxes of Zn into the root stele (and its effects on Zn efflux from the stele) remain unclear, as does the distribution of this character among closely related taxa. Further anatomical studies in the Thlaspiceras/Noccaea/Raparia/Microthlaspi/Neurotropis clade are likely to be revealing in this respect. In a closely related Brassicaceae species, Thellungiella halophila O. E. Schulz, a second layer of endodermis is developed and it is probably related to the salt tolerance of this halophyte (Inan et al., 2004). Thus, the structural/functional adaptations of roots associated with metallophily are highly variable in this interesting group of plants.

Figure 2.

Development scheme of apoplastic barriers along the root axis of Thlaspi caerulescens (hypertolerant Zn-accumulator) and T. arvense (nontolerant Zn nonaccumulator). In the first stage of endodermal ontogenesis, Casparian bands, and in the second stage, suberin lamellae deposition, develop closer to the root tip in T. caerulescens than in T. arvense. Note the early formation of a peri-endodermal layer close to the apex in T. caerulescens but not in T. arvense. Bar, 2 mm in the longitudinal axis. Transverse axes are not represented to scale.

Figure 3.

Structure of primary roots in cross-sections of Thlaspi arvense (a) and T. caerulescens (b–f). (a) Autofluorescence of T. arvense root section showing typical dicotyledonous root structure with three cortical layers; the innermost is the endodermis exhibiting fluorescence of Casparian bands. (b) Autofluorescence of T. caerulescens root section showing the intensive fluorescence of inner tangential and radial walls of the peri-endodermal layer and light fluorescence of thin endodermal cell walls. (c) T. caerulescens primary root with emerging lateral root primordium in fluorescence microscope. After Fluorol yellow-toluidine blue staining, endodermal cell walls exhibit intensive bright yellow fluorescence as a result of suberin lamellae deposition (second stage of endodermal development). (d) Semithin section of Spurr embedded T. caerulescens root showing prominent wall ingrowths of the peri-endodermal layer (toluidine blue-basic fuchsin staining). (e) Transmission electron microscopy (TEM) of the young root part of T. caerulescens (< 1 mm from the apex) stained by KMnO4 to visualize Casparian bands, which are present in radial walls in endodermis, but absent in the peri-endodermal layer. (f) TEM of the older root part of T. caerulescens (c. 2.5 mm from the apex) with irregular wall thickening in the peri-endodermal layer. Cb, Casparian band; e, endodermis; ic, intercellular space; p, pericycle; p-e, peri-endodermal layer; arrow, wall ingrowths in peri-endodermal cells. Bars, 50 µm (a–d); 4 µm (e, f).

Molecular insights into Zn hyperaccumulation  Gene and protein expression profiling (Assunção et al., 2001; Becher et al., 2004; Weber et al., 2004; Craciun et al., 2006; Filatov et al., 2006; Hammond et al., 2006; Rigola et al., 2006; Talke et al., 2006; Tuomainen et al., 2006; van de Mortel et al., 2006; Weber et al., 2006), and functional analyses of proteins in heterologous plant, yeast and Xenopus laevis oocyte systems are providing remarkable and rapidly advancing insights into Zn hyperaccumulation – and Zn homeostasis in general – at the molecular level. Fortunately, T. caerulescens and A. thaliana share > 87% sequence identity (Peer et al., 2003; Rigola et al., 2006), enabling A. thaliana sequence data to be easily exploited. To illustrate, global shoot transcriptome data published using the full-genome Affymetrix A. thaliana ATH1-121501 (ATH1) GeneChip array (Affymetrix Inc. Palo Alto, CA, USA), for T. caerulescens vs T. arvense (Hammond et al., 2006), and A. halleri vs A. lyrata ssp. petraea (Filatov et al., 2006) and A. thaliana (Talke et al., 2006) comparisons, are combined and re-analysed here. Briefly, raw expression data files (CEL files) of shoot transcriptome data from either the hyperaccumulator (five T. caerulescens, eight A. halleri) or nonhyperaccumulator (five T. arvense, six A. lyrata ssp. petraea, two A. thaliana) condition, were analysed using a global Robust Multichip Average (RMA) algorithm (Irizarry et al., 2003), with a custom GeneChip definition file (CDF) in GeneSpring GX (Agilent Technologies Inc.). This CDF was designed to exclude ATH1 GeneChip array oligonucleotide probe pairs that hybridized to both T. caerulescens and T. arvense genomic DNA (gDNA) below an intensity threshold of 300 (sensu Hammond et al., 2005, 2006). Thus, 22 131 probe sets (out of 22 746), most likely to be common to Thlaspi s.l and Arabidopsis, were used for transcriptome comparisons. Data were normalized to the median expression value of either the hyperaccumulator or the nonhyperaccumulator condition. Putative genes with different amounts of expression between the two conditions were identified using a Welch's t-test with a Benjamini-Hochberg False Discovery Rate (FDR) correction of 0.05 (Table S10).

In total, homologues of 60 A. thaliana genes are significantly differentially expressed between hyperaccumulators and nonhyperaccumulators and may have conserved roles in brassicaceous Zn hyperaccumulation (Table S10). Six of these genes encode proteins with putative roles in Zn transport: three cation diffusion facilitator (CDF) family members (At2g39450 (TAIR6: AtMTP11), At2g46800 (AtZAT1/AtMTP1) and At3g58060 (AtMTPc3)); a member of the Zn-Fe permease (ZIP) family (At1g60960 (AtIRT3/TcZNT2)); and a PIB-type heavy-metal-associated domain-containing ATPase (At4g30120 (AtHMA3)). CDFs appear to mediate vacuolar sequestration of Zn through Zn efflux from the cytoplasm (van der Zaal et al., 1999; Persans et al., 2001; Blaudez et al., 2003; Delhaize et al., 2003; Hall & Williams, 2003; Dräger et al., 2004; Kim et al., 2004; Kobae et al., 2004; Desbrosses-Fonrouge et al., 2005; Krämer, 2005b; Arrivault et al., 2006; Colangelo & Guerinot, 2006), ZIPs are likely to mediate cellular Zn uptake (Pence et al., 2000; Assunção et al., 2001; López-Millán et al., 2004; Colangelo & Guerinot, 2006), and HMAs have Zn-transport functions throughout the cell (Williams et al., 2000; Mills et al., 2003; Bernard et al., 2004; Eren & Argüello, 2004; Gravot et al., 2004; Hussain et al., 2004; Papoyan & Kochian, 2004; Verret et al., 2004, 2005; Williams & Mills, 2005; Colangelo & Guerinot, 2006). However, the function of other nutritionally important transporter genes, for example, a phosphate-starvation-inducible high-affinity phosphate transporter AtPT2/AtPHT4/AtPHT1;4 (Misson et al., 2004), which is highly expressed in Zn hyperaccumulators (Hammond et al., 2006; Talke et al., 2006), remains unknown. Other genes expressed highly in Zn hyperaccumulators include plant defensins (PDFs), which confer Zn tolerance and accumulation in heterologous systems, and may act as blockers of Zn-permeable channels (Mirouze et al., 2006). Whilst it remains extremely challenging to elucidate how the function of single proteins in heterologous systems relates to the behaviour at whole-plant and crop scales, the use of homologous transformation (Peer et al., 2003) and the increasing availability of species-specific sequence information (Rigola et al., 2006) will undoubtedly hasten this process.

Acknowledgements

We apologize to those authors whose work has not been cited owing to space constraint or oversight. We especially thank R. D. Reeves (Palmerston North) for kindly providing unpublished data and stimulating discussion during the drafting of the table of Zn accumulators (Table 3). We thank friends and colleagues for their insightful comments and discussions on early drafts, including A. J. M. Baker (Melbourne), N. Verbruggen (Brussels), S. N. Whiting (Golders), M. Wissuwa (Tsukuba) and S. D. Young (Nottingham). We thank U. Krämer and I. N. Talke (Potsdam-Golm) for providing CEL files for our analyses. All CEL files are available from NASC (http://arabidopsis.info/). I. Zelko and A. Lux gratefully acknowledge Z. Šulavíková for technical support, I. M. Čaplovičová for assistance with the TEM, and the Slovak Ministry of Education for financial support to COST 859, APVV 51-013304 and VEGA 1-4354-07 projects.

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