A mutation in the Arabidopsis thaliana cell wall biosynthesis gene pectin methylesterase 3 as well as its aberrant expression cause hypersensitivity specifically to Zn


For correspondence (e-mail stephan.clemens@uni-bayreuth.de).


Defects in metal homeostasis factors are often accompanied by the loss of metal tolerance. Therefore, we screened for mutants with compromised growth in the presence of excess Zn2+ in order to identify factors involved in Zn biology in plants. Here we report the isolation of six ozs (overly Zn sensitive) ethyl methanesulfonate Arabidopsis thaliana mutants with contrasting patterns of metal sensitivity, and the molecular characterization of two mutants hypersensitive specifically to Zn2+. Mutant ozs1 represents a non-functional allele of the vacuolar Zn transporter AtMTP1, providing additional genetic evidence for its major role in Zn2+ tolerance in seedlings. Mutant ozs2 carries a semi-dominant mutation in the gene encoding pectin methylesterase 3 (AtPME3), an enzyme catalyzing demethylesterification of pectin. The mutation results in impaired proteolytic processing of AtPME3. Ectopic expression of AtPME3 causes strong Zn2+ hypersensitivity that is tightly correlated with transcript abundance. Together these observations suggest detrimental effects on Golgi-localized processes. The ozs2 but not the ozs1 phenotype can be suppressed by extra Ca2+, indicating changes in apoplastic cation-binding capacity. However, we did not detect any changes in bulk metal-binding capacity, overall pectin methylesterification status or cell wall ultrastructure in ozs2, leading us to hypothesize that the ozs2 mutation causes hypersensitivity towards the specific interference of Zn ions with cell wall-controlled growth processes.


The bioavailability of the micronutrient Zn was recently proposed as a limiting factor in the evolution of eukaryotes (Dupont et al., 2010). Eukaryotic diversification coincided with the transition to an oxidizing ocean, concomitant higher Zn solubility under these geochemical conditions and a marked expansion of Zn usage about 800–500 million years ago. Eight to 10% of the proteins in modern eukaryotes are estimated to be Zn-binding compared with 5–6% for bacteria and archaea (Andreini et al., 2006), making Zn the most widely recruited microelement in eukaryotes. Zn-dependent proteins are found in all six Enzyme Commission (EC) classes, most prominently among hydrolases (EC3) (Andreini et al., 2008). The second major function of Zn in biology besides catalysis lies in the stabilization of protein structures. In fact, the higher proportion of Zn-binding proteins in eukaryotic proteomes is largely due to the recruitment of Zn for structural roles such as in Zn-finger domain proteins.

Zn(II) strongly interacts with N-, O- and S-containing organic molecules, for instance histidines, aspartic or glutamic acids, and cysteines, the major Zn(II) ligands in proteins. It can replace other essential cations in biological molecules according to the Irving–Williams series and thus become toxic to cells when present in excess. Zn toxicity is not only a consequence of soil metal contamination through industrial activities such as mining and metal smelting (Chaney, 2010). It can also occur under conditions such as Fe deficiency when Zn availability and influx are high relative to those of Fe and can result in displacement of Fe from Fe-binding sites in proteins (Arrivault et al., 2006). Homeostatic mechanisms ensure uptake, distribution as well as storage of Zn, and suppress uncontrolled interaction of Zn with proteins (Palmer and Guerinot, 2009; Clemens, 2010; Sinclair and Krämer, 2012). In spite of the vastly fluctuating availability of Zn in the soil, the concentration in plant tissues is usually maintained at 15–50 μg g−1 dry weight (DW) (Hänsch and Mendel, 2009). Integral components of Zn homeostasis are Zn transporters and Zn-chelating molecules.

An essential aspect of metal homeostasis is basal metal tolerance. It can be regarded as a consequence of homeostatic mechanisms. Many components of metal distribution and buffering are key tolerance factors (Krämer and Clemens, 2005). Their loss of function results in inhibition of growth under conditions of supraoptimal metal availability. In the case of Zn, well-documented examples are heavy metal ATPases (HMAs), metal tolerance proteins (MTPs) and zinc-induced facilitator 1 (ZIF1). Arabidopsis thaliana HMA2 and HMA4 as well as Arabidopsis halleri HMA4 are not only essential for the translocation of Zn from the root to the shoot (Hussain et al., 2004) but also confer Zn tolerance, as the efflux activity in root cells not surrounding the vasculature reduces cytosolic Zn overload (Verret et al., 2004; Mills et al., 2005; Hanikenne et al., 2008). Similarly, a defect in the Zn-efflux system PCR2 causes Zn hypersensitivity as well as a reduction in Zn translocation (Song et al., 2010). Metal tolerance proteins and other transporters such as ZIF1 are important for storage and detoxification of Zn (Kobae et al., 2004; Desbrosses-Fonrouge et al., 2005; Arrivault et al., 2006; Haydon and Cobbett, 2007; Haydon et al., 2012). The cytosolic thiol Zn chelators glutathione and phytochelatins contribute to tolerance too (Tennstedt et al., 2009; Shanmugam et al., 2012). Thus, a suitable approach for identifying novel components of Zn homeostasis appears to be the screening for Zn-hypersensitive mutants. We therefore performed a screen for ethyl methanesulfonate (EMS) mutants impaired in their ability to grow in the presence of toxic concentrations of Zn2+. Six mutants with contrasting metal specificities were identified. Their isolation and the molecular characterization of two specifically Zn2+-hypersensitive mutants are described here. One of the mutants represents a new allele of MTP1 while the other carries a mutation in pectin methylesterase 3 (AtPME3, At3g14310).

Binding of metal ions to the cell wall has long been discussed as a mechanism of metal tolerance, although direct evidence is limited (Hall, 2002). Many studies on plants exposed to excess metal have detected metal immobilization in the cell wall (Krzeslowska, 2011). Most important for the binding capacity are homogalacturonans, one of the polysaccharide domains of pectin (Caffall and Mohnen, 2009). Pectins are the most complex wall components (Wolf and Greiner, 2012). They are secreted by plant cells in a highly methylesterified form. Pectin methylesterases (PMEs; EC are responsible for the spatially regulated de-esterification of pectin in the cell wall. Through this activity PMEs can generate free carboxylic groups which can interact with Ca2+ ions. Because these Ca2+-binding sites of demethylated pectin account for most of the cation exchange capacity of cell walls, pectin modulation could affect the metal immobilization capacity of the cell wall and thereby the metal ion activity near the plasma membrane. However, we found no evidence for alterations of bulk cell wall metal binding in our mutant. Instead, the effects of the isolated AtPME3 mutation as well as constitutive overexpression of AtPME3 indicate an interference of Zn with cell wall-controlled growth processes.


Identification of Zn-hypersensitive A. thaliana mutants

With the aim of identifying new components of the plant Zn homeostasis network, we screened EMS mutagenized M2 seedlings for reduced growth under conditions of excess Zn2+ on dilute Hoagland medium found previously to allow more sensitive detection of Zn2+ tolerance phenotypes (Tennstedt et al., 2009). About 24 000 seedlings were cultivated in the presence of 60 μm Zn2+, a concentration that inhibited Col-0 wild-type seedlings under these conditions by about 40%. Seedlings showing reduced root length compared with Col-0 wild type were transferred to plates with control Zn2+ concentration and recovery of root elongation was scored (Figure S1 in Supporting Information). In order to eliminate seedlings with general growth defects, only seedlings displaying growth similar to co-transferred wild-type seedlings were selected. A total of 359 putative mutants were isolated. The progeny of these plants were tested again for hypersensitivity to Zn2+. Finally, six lines were confirmed to show significantly stronger root growth inhibition than Col-0. They were established as ozs (overly Zn sensitive) mutants.

Based on root growth rate under control conditions we divided the mutants into two groups: group one consisting of four mutants (ozs1, ozs2, ozs5 and ozs6) showing wild-type growth, and group two consisting of two mutants (ozs3 and ozs4) showing a significant reduction compared with Col-0 (Figure 1). F1 phenotypes of crosses with Col-0 indicated that mutants ozs1, ozs3, ozs4, ozs5 and ozs6 are recessive, while ozs2 is semi-dominant. Complementation analysis revealed that the six lines carry mutations in six different loci.

Figure 1.

Zn2+-hypersensitivity of six isolated ozs mutants. Wild type (Col-0) and ozs mutants were grown vertically under control conditions or in the presence of 60 μm ZnSO4. The length of the primary roots was determined after about 12 days of cultivation (when Col-0 seedlings on control plates had reached the bottom of the plate). Data represent three to five independent experiments (= 77–102). (a) Mean values ± SD. Significant differences from wild type were determined by two-way anova and the Tukey test, *P < 0.05, **P < 0.01, ***P < 0.001. Mutants which show no significant reduction in root length under control conditions were placed in group 1, mutants with reduced root length under control conditions in group 2. (b) Growth under control condition was set to 100% and relative root growth at 60 μm ZnSO4 was calculated. Significant differences from wild type were determined by one-way anova and the Tukey test. ***P < 0.001.

Next we performed specificity screening by testing the mutants for hypersensitivity towards other metal cations. Three mutants (ozs1, ozs2, ozs3) showed a highly significant increase in sensitivity exclusively towards Zn2 + . Mutant ozs4 was more sensitive towards Zn2+, Cd2+, Cu2+, Ni2+ and Co2+ compared with the Col-0 wild type. Mutant ozs5 displayed an enhanced sensitivity towards Zn2+ and Cu2+. Zn2+, Cd2+, Cu2+ and Mn2+ treatment caused a stronger reduction in root growth in ozs6 as compared with the wild type (Table 1). These findings indicated that different mechanisms contributing to Zn2+ tolerance are affected in the isolated mutants.

Table 1. Specificity of metal hypersensitivity of the ozs mutants. All identified mutants were grown in the presence of different metal excess conditions to assay metal hypersensitivity relative to Col-0. Concentrations of the different metals (60 μm Zn2+, 2 μm Cd2+, 10 μm Cu2+, 300 μm Fe2+, 6 μm Co2+, 450 μm Mn2+, 6 μm Ni2+) were adjusted in a way that all treatments had similar impact on Col-0 (growth reduction of 50–60% compared with control conditions). Relative root growth rates were determined by cultivating seedlings on control plates and on plates containing the respective metal cation excess. Data represent three to six independent experiments, n = 43–107 seedlings. Significant differences from Col-0 in metal sensitivity were determined by two-way anova and the Tukey test
  1. a

    P < 0.05.

  2. b

    P < 0.01.

  3. c

    P < 0.001.

ozs1 c
ozs2 c
ozs3 c
ozs4 c c c a c
ozs5 c b
ozs6 c c c c

ozs1 is a loss-of-function allele of AtMTP1

We first studied ozs1 and ozs2, the two Zn2+-specific mutants with the strongest hypersensitivity and wild-type growth under control conditions. Two backcross lines each were isolated and used for characterizing the mutant phenotypes. Hypersensitivity of ozs1 is already apparent at very low concentrations of 40 μm Zn2+ (Figure S2). Mutant ozs1 was crossed to Ler wild type to generate a segregating F2 population. Ler shows a growth response to excess Zn2+ similar to that of Col-0. In this mapping population we observed a ratio of approximately 3:1 of wild-type to Zn2+ hypersensitivity phenotypes, indicating a monogenic recessive trait. The ozs1 mutation was mapped in about 400 phenotyped F2 plants to a 279-kb interval on the lower arm of chromosome 2. One prominent candidate gene (AtMTP1, At2 g46800) is located in this area and was therefore sequenced in ozs1. We found one nucleotide exchange from G to A, which leads to an amino acid change at position 293 from aspartic acid to asparagine. This renders the protein non-functional, as shown by heterologous expression of the wild-type and ozs1 version in the Saccharomyces cerevisiae mutant zrc1cot1 (Figure S3a,b). A wild-type genomic fragment of AtMTP1 complemented the phenotype (Figure S3c). MTP1 mediates Zn transport into the vacuole and is the best-characterized Zn tolerance factor in A. thaliana (Kobae et al., 2004; Desbrosses-Fonrouge et al., 2005). Thus, we interpreted the finding that a mutation in MTP1 is causing the ozs1 phenotype as validation of our genetic screen and focused further studies on ozs2.

Physiological characterization of ozs2

First we tested hypersensitivity at different Zn2+ doses. Significantly stronger growth reduction relative to wild type was observed for ozs2 seedlings at Zn2+ concentrations ≥60 μm (Figure 2). Next we addressed the specificity of the hypersensitivity. In addition to other metal cations we tested the effects of salt, osmotic and oxidative stress on root growth. Seedlings were exposed to concentrations of the respective stress factors that caused a reduction of about 50% in root elongation in Col-0. None of the tested conditions revealed any differences between ozs2 and the wild type, indicating Zn2+-specific hypersensitivity (Figure S4). Microscopic analysis showed that the Zn2+-dependent inhibition of root elongation is largely attributable to a defect in cell elongation. The root cell size of seedlings grown in the presence of 80 μm Zn2+ was about 40% smaller in ozs2 than in Col-0, while no difference was observed under control conditions (Figure 3c,d). Morphological changes in ozs2 under excess Zn2+ include, beside strong reduction in the size of the cell elongation zone, an increase in the number of root hairs relative to Col-0 (Figure 3a,b).

Figure 2.

Dose-dependent reduction in ozs2 root growth under excess Zn2+. Seedlings of Col-0 and two independent ozs2 backcross lines (BC) were grown vertically in the presence of different concentrations of ZnSO4. The length of the primary roots was determined after about 12 days of cultivation (when Col-0 seedlings on control plates had reached the bottom of the plate). (a) Data representing mean values ±SD of three independent experiments (n = 45–75). Significant differences from wild type were determined by two way anova and the Tukey test. ***P < 0.001. (b) Seedlings after 12 days' cultivation, bar = 1 cm.

Figure 3.

Reduction in root cell size and stimulation of root hair formation in ozs2 seedlings exposed to excess Zn2+. Col-0 and ozs2 seedlings were grown for 5 days under control conditions or in the presence of 80 μm ZnSO4. (a) Root morphology, bar = 100 μm. (b) The number of root hairs was determined in the first millimeter of the differentiation zone. Mean values ± SD for two independent experiments are shown (n = 7–20). (c) Seedlings were stained with propidium iodide and analyzed by confocal laser microscopy, bar = 50 μm. (d) Cell size was determined using ImageJ. Data represent means ± SD of three independent experiments (five roots each). Significant differences from wild type were determined by Student's t-test. ***P < 0.001.

Possible effects of the ozs2 mutation on Zn accumulation in roots and shoots were tested in the vertical plate system used for tolerance assays. Elemental profiles were obtained for Col-0 and ozs2 seedlings cultivated under control and excess Zn2+ conditions. No significant differences in Zn accumulation were detected on control medium or in the presence of 40 μm Zn2+. When assayed at a concentration of 60 μm Zn2+, which inhibits growth of ozs2 seedlings more strongly than Col-0, slightly higher Zn accumulation was observed in ozs2 shoots. However, Zn accumulation in roots, i.e. the organs that show the hypersensitivity phenotype, was not significantly different between the genotypes even under these conditions (Figure S5).

Genetic characterization and map-based cloning of ozs2

Root growth assays with the F1 generation of a cross between ozs2 and Col-0 indicated, that the mutation in ozs2 has a semi-dominant effect at 80 μm Zn2+ (Figure S6). The reduction of root growth was intermediate between Col-0 and the mutant. For the F2 generation a ratio of 164:285:149 (wild type:intermediate sensitivity:full hypersensitivity) was obtained, consistent with semi-dominance (χ2 = 2.06, P = 0.356). To identify the gene responsible for the hypersensitivity of ozs2 we again followed a map-based cloning approach. F2 plants showing strong Zn2+ hypersensitivity were selected from a cross of ozs2 with Ler. The ozs2 mutation was localized to the upper arm of chromosome 3 and further linkage analyses defined a 54-kb region which included eight open reading frames. All predicted genes were sequenced in ozs2. The only mutation found was a nucleotide exchange from T to C in the third exon of the At3g14310 gene. This exchange leads to a substitution of glycine to valine. The At3g14310 gene encodes pectin methylesterase 3, which catalyzes the demethylesterification of cell wall pectin (Micheli, 2001). At3g14310 belongs to a 66-member multigene family in A. thaliana (Pelloux et al., 2007). The encoded protein consists of 592 amino acids and has a molecular weight of 64.3 kDa. The detected amino acid exchange is located within a five amino acid stretch which is highly conserved in PMEs from different plants, fungi and bacteria (Figure 4).

Figure 4.

The ozs2 version of AtPME3 carries a mutation in a highly conserved amino acid. Partial amino acid sequence alignment for pectin methylesterases from higher plants, fungi, and bacteria: AtPME3 (Arabidopsis thaliana; O49006), Nicotiana tabacum (Q9LEB0), Medicago truncatula (Q9SC89), Petunia inflata (Q43043), Solanum tuberosum (Q9SEE6), Pyrus communis (A0ZNK1), Vitis vinifera (F6HXK8), Coffea arabica (I1UYB4), Oryza sativa (japonica) (Q6ZDX2), Zea mays (O24596), Aspergillus niger (P17872), Botrytis cinerea (Q9C2Y1), Clostridium acetobutylicum (Q97DU8), Yersinia enterocolitica (A1JJ76). The arrow marks the amino acid that is mutated in ozs2 to Val. Conserved residues are highlighted in black.

However, the apparent semi-dominant character of ozs2 had already suggested that the mutant phenotype is not caused by a loss of AtPME3 function. Correspondingly, two independent homozygous AtPME3 GABI-Kat lines (002A10 and 329D007) (Figure S7) with T-DNA insertions in exon 1 and exon 3, respectively, grew like Col-0 on medium with toxic concentrations of Zn2+. We therefore attempted rescue of the ozs2 mutant by suppressing AtPME3 expression through a RNA interference (RNAi) approach and generated several transgenic lines in ozs2 background with a strongly reduced AtPME3 transcript abundance. AtPME3 transcript levels in untransformed ozs2 plants were undistinguishable from wild type. We chose two lines with the strongest RNAi effect for further analysis (Figure 5a). Both lines showed a complete rescue of the ozs2 mutant phenotype (Figure 5b,c). Taken together, these data strongly suggested that the mutation in AtPME3 is semi-dominant and indeed responsible for the Zn2+ hypersensitivity of ozs2.

Figure 5.

Rescue of the mutant phenotype by RNA interference (RNAi)-mediated knock-down of AtPME3 in ozs2. (a) Transcript abundance of AtPME3 was determined by quantitative real-time PCR in Col-0, ozs2, and two independent homozygous AtPME3-RNAi lines in ozs2 background (A3 and C1), grown in liquid culture to comparable size (7–9 days). The ΔCT value was calculated by subtracting the cycle threshold (CT) value of the reference gene EF1α from the CT value of the target gene PME3. The relative transcript level (RTL) was calculated as: RTL = 1000 × 2−ΔCT. Data represent means of three independent experiments. (b) Col-0, ozs2, and the two AtPME3-RNAi lines A3 and C1 were grown vertically for 12 days in the presence of 80 μm ZnSO4, bar = 1 cm. (c) The length of the primary root was determined after about 12 days of cultivation (when Col-0 seedlings on control plates had reached the bottom of the plate). Data represent means ± SD of three independent experiments (= 31–68). Significant differences from wild type were determined by two-way anova and the Tukey test. ***< 0.001.

Overexpression of AtPME3 leads to strongly enhanced Zn2+ hypersensitivity

To further support this hypothesis we overexpressed the ozs2 mutant version of AtPME3 (AtPME3-mut) in A. thaliana Col-0. Several strongly expressing lines were isolated that showed a far more severe Zn2+ hypersensitivity than the original ozs2 mutant (Figures 6a and S8). Growth was inhibited by about 80% on medium containing 40 μm ZnSO4, a concentration that inhibited ozs2 seedlings by only about 40% (Figure 2). Again, this phenotype appeared to be Zn2+-specific as no growth reduction stronger than for Col-0 was observed in the presence of toxic concentrations of either Cd2+ or Co2+ (Figure S8b). Surprisingly, control experiments revealed that lines overexpressing wild-type AtPME3-GFP in the Col-0 background were nearly as hypersensitive to Zn2+ as plants overexpressing AtPME3-mut-GFP (Figure 6a). These effects were not caused by the GFP fusion, as lines overexpressing the two different AtPME3 versions without GFP displayed very similar phenotypes.

Figure 6.

Overexpression of wild-type (WT) AtPME3, the ozs2 AtPME3, and an enzymatically inactive version of ozs2 AtPME3 resulted in strong Zn2+ hypersensitivity. (a), (b) Seedlings of Col-0 and various homozygous transgenic lines in the Col-0 background overexpressing either the wild type AtPME3 (=PME3-OX), the ozs2 mutant version (=PME3-mut-OX) (a), or an enzymatically inactive version of AtPME3-mut (=PME3-mut-ia-OX1 and PME3-mut-ia-OX2) (b) were grown vertically under control conditions and in the presence of 40 μm ZnSO4. The length of the primary root was determined after about 12 days of cultivation (when Col-0 seedlings on control plates had reached the bottom of the plate). Data represent means ± SD of three independent experiments (= 30–72 for (a), = 34–51 for (b)). Significant differences from wild type were determined by two-way anova and the Tukey test. ***< 0.001. Please note that transcript data for the overexpression lines used in (a) and (b) are shown in Figure S8a. (c), (d) Transgenic lines overexpressing either the wild-type AtPME3 version (c) or the ozs2 mutant version (d) in the atpme3 T-DNA insertion line were tested for AtPME3 and AtPME3-mut transcript levels, respectively, and inhibition of root growth in the presence of 40 μm ZnSO4. Growth reduction was calculated based on the mean values for primary root length under control and excess Zn2+ conditions (two or three independent experiments, = 14–16 per experiment). Transcript levels, expressed relative to EF1α, were determined by quantitative real-time PCR for seedlings grown in liquid culture to comparable size (7–9 days).

In order to compare the overexpression effects of wild-type and ozs2 mutant PME3 more closely we generated additional transgenic lines. We chose atpme3 KO plants (Gabi-Kat 002A10) as the genetic background to exclude any effects possibly caused by an interaction of wild-type and ozs2 mutant AtPME3. We analyzed 10 or more lines for AtPME3 and AtPME3-mut transcript levels, respectively, and observed a direct correlation between the transcript level and the degree of Zn2+ hypersensitivity (Figure 6c,d). Regardless of the AtPME3 version overexpressed, higher transcript abundance resulted in stronger growth inhibition by excess Zn2+. No difference between wild-type and mutant AtPME3 was observed in the impact of comparable transcript levels on growth at 40 μm Zn2+.

Next we tested whether the semi-dominant effect of the ozs2 mutation is dependent on PME enzyme activity. Similar to Dorokhov et al. (2006) we substituted amino acids 408 (glutamine) and 409 (asparagine) with alanine. These amino acids correspond to positions 135 and 136 in carrot PME for which the structure was solved (Johansson et al., 2002). They are part of the active center and essential for activity. The resulting dead version was overexpressed in atpme3. Again, we found pronounced Zn2+ hypersensitivity in all strongly expressing plants (Figure 6b), suggesting that the ozs2 phenotype is not caused by a change in catalytic activity of PME3. Total PME activity was tested using three different assays, at varying pHs ranging from 5.0 to 8.0 and with citrus pectins of different degrees of methylesterification as substrates. No difference could be detected between wild-type plants, atpme3 and lines strongly overexpressing either the Col-0 or the ozs2 version of PME3 in the atpme3 background with any of the assays. Figure S9 shows the results obtained with the methanol oxidation assay according to Reca et al. (2012).

The ozs2 mutation causes a PME processing defect

Type-I PMEs such as PME3 undergo proteolytic processing. The protein contains a pro-region that is cleaved during protein maturation (Pelloux et al., 2007). The two conserved cleavage motifs are discernible in AtPME3 (Figure 7a). We tested whether the ozs2 mutation affects processing by expressing hemagglutinin (HA)-tagged versions of wild-type and ozs2 mutant PME3 in A. thaliana. Western blot analysis of seedling protein extracts revealed three products. Their sizes corresponded to the unprocessed protein and two cleavage products (Figure 7b). This pattern was comparable to the one previously reported for a HA-tagged Nicotiana tabacum PME transiently expressed in N. benthamiana (Wolf et al., 2009). Interestingly, the relative abundance of the three proteins was consistently different for the AtPME3 carrying the ozs2 mutation, as far more of the pro-protein remained unprocessed (Figure 7c). This difference between wild-type and mutant protein was unaffected by exposure to Zn2+. Thus, the ozs2 mutation appears to affect PME3 processing. This and the strong phenotypes caused by PME3 overexpression suggest that Zn2+ hypersensitivity could be due to an interference of mutated or aberrantly expressed AtPME3 with processes involved in the synthesis and/or modification of the cell wall. Therefore, we asked whether the ozs2 mutation causes changes in the cell wall that could explain the observed phenotype.

Figure 7.

Impaired proteolytic processing of the ozs2 mutant version of AtPME3 pro-protein. The pro-region of type-I pectin methylesterase (PMEs) such as AtPME3 is proteolytically removed. (a) A diagram representing the AtPME3 domain structure according to Wolf et al. (2009). Indicated are signal peptide (SP), pro-region, the two basic cleavage motifs, the PME domain, and the C-terminal hemagglutinin (HA) tag used for immunodetection. (b) Seedlings overexpressing either the Col-0 or the ozs2 mutant version of AtPME3-HA in the Col-0 background were grown vertically on control and excess Zn2+ plates (50 μm). After 10 days seedlings were harvested and protein extracted. Western blots were stained with an anti-HA monoclonal antibody. (c) ImageJ quantification of signals corresponding to the pro-protein and the two processed proteins. Shown are mean values ± SD of three independent experiments.

Excess Ca2+ suppresses the Zn2+ hypersensitivity of ozs2 but not of ozs1

Demethylesterified pectin is known to form Ca2+-binding sites (Pelloux et al., 2007). They can potentially be occupied by other cations as well and thereby influence metal tolerance (Hall, 2002). For that reason we tested the effect of excess Ca2+ on the Zn2+ hypersensitivity of ozs2 seedlings in comparison with ozs1 as a mutant deficient in vacuolar sequestration of Zn, and hence presumably with unaffected cell wall architecture. Seedlings of mutant backcross lines were exposed to excess Zn2+ that resulted in similar degrees of inhibition of root growth, i.e. 40 μm for ozs1 and 60 μm for ozs2. Hypersensitivity of ozs1 was not influenced by the addition of 0.5 mm extra Ca2+ (Figure 8). In contrast, the phenotype of ozs2 seedlings was completely suppressed in high-Ca2+ medium. The same effect was found for seedlings overexpressing PME3 in atpme3 (Figure S10). These results indicated that changes in Ca2+-binding sites of the cell wall could be underlying the ozs2 mutant phenotype. We thus searched for possible ozs2-dependent alterations in cell wall architecture.

Figure 8.

Suppression of the ozs2 but not the ozs1 Zn2+-hypersensitivity phenotype by high Ca2+. Zn2+ tolerance of Col-0 and two independent backcross lines of ozs2 (a) and ozs1 (b) each was assayed either in the absence or the presence of 0.5 mm CaCl2. Seedlings were exposed to 60 μm for ozs2 and to 40 μm for ozs1 in order to elicit similar degrees of growth inhibition. The length of the primary roots was determined after about 12 days of cultivation (when Col-0 seedlings on control plates had reached the bottom of the plate). Data represent means ± SD of three independent experiments (= 43–48). Significant differences from Col-0 were determined by two-way anova and the Tukey test. *< 0.05; ***< 0.001.

Microscopic analysis of cell wall structure in ozs2

Monoclonal antibodies can be used to visualize changes in the degree of methylesterification or pattern of pectin (Lee et al., 2011). We stained sections of Col-0 and ozs2 roots with JIM7 and CCRC-M38. Epitopes recognized by these antibodies are heavily methylesterified homogalacturonan and de-esterified homogalacturonan, respectively. Roots were analyzed at various distances from the tip. The pattern and abundance of JIM7 or CCRC-M38 epitopes change during root development in Col-0 as well as in ozs2. However, no consistent differences in staining intensity could be detected. Quantification of signals did not yield significant differences (Figure S11), indicating that the ozs2 mutation does not result in robust changes in methylesterification relative to Col-0.

In addition, we analyzed the root ultrastructure of control and ZnSO4-treated Col-0 and ozs2 seedlings by electron microscopy. Cross-sections did not reveal any obvious dissimilarities between the genotypes. Cell wall thickness was determined for all root cell types. While there was variation apparent, no consistent differences separating Col-0 from the two backcross lines were detected (Table S1). Moreover, no ozs2-dependent ultrastructural changes were apparent in the stele where AtPME3 transcript is most abundant (Guénin et al., 2011) (Figure S12). Because PMEs are trafficked to the cell wall through the secretory pathway, the Golgi was analyzed in more detail. However, no alterations in size, appearance or localization of Golgi stacks were apparent in the two investigated ozs2 backcross lines. We concluded that the ozs2 mutation does not affect cell wall architecture in a manner that is detectable by microscopic techniques.


Few factors have been identified to date that help plants tolerate toxic conditions of excess Zn2+, which can occur, for instance, as a consequence of mining, metal smelting, or application of sewage sludge to agricultural soil. Understanding metal tolerance is a prerequisite for biotechnological approaches to phytoremediation (Sinclair and Krämer, 2012). Perhaps even more importantly, basal metal tolerance is a consequence of mechanisms of metal homeostasis. Exploration and dissection of mechanisms of Zn2+ tolerance can therefore advance the understanding of the interplay of Zn transport and Zn-binding activities that result in the safe trafficking of Zn to the large number of target sites in tissues, cell types and cellular compartments of a plant. Also, it can help uncover novel aspects of the biological roles played by Zn. In spite of this potential, Zn2+ tolerance is as yet genetically underexplored (Richard et al., 2011).

The specificities of the isolated ozs mutants can help unravel metal homeostasis and metal toxicity

Known Zn2+ tolerance mechanisms differ in their metal specificity. While vacuolar sequestration in A. thaliana is mediated by the Zn-specific transporters MTP1 and MTP3 (Kobae et al., 2004; Desbrosses-Fonrouge et al., 2005; Arrivault et al., 2006), efflux is dependent on Zn- and Cd-transporting P1b-ATPases such as HMA2 and HMA4 (Hussain et al., 2004; Mills et al., 2005). Similarly, cytosolic Zn buffering is partly dependent on phytochelatins, which efficiently bind Cd as well (Tennstedt et al., 2009). The results of our genetic screen revealed different sensitivity patterns too. Mutants ozs1, ozs2 and ozs3 are hypersensitive specifically to excess Zn2+, while ozs4, ozs5 and ozs6 also show growth inhibition in the presence of a varying set of other metal cations. The observed combinations do not immediately suggest the involvement of any known metal tolerance pathways. All three mutants share Cu2+ hypersensitivity. Two of them show Cd2+ hypersensitivity as well. Taken together, these findings suggest the existence of both a variety of Zn toxicity targets and Zn tolerance mechanisms that can now be elucidated through the molecular characterization of the ozs mutants.

Furthermore, the Zn2+-specific hypersensitivity of ozs1 and ozs2 had already demonstrated that the mutant phenotypes are not the result of a general higher susceptibility to oxidative stress, which is often a consequence of exposure to toxic metals (Clemens, 2006). This was further supported by the wild-type growth of ozs2 when exposed to other abiotic stresses (Figure S4). Thus, the ozs mutants are likely to be affected in processes that are directly linked to metal biology.

ozs1 represents a loss-of-function allele of AtMTP1

Mapping of the causal mutation in ozs1 led to the identification of a new atmtp1 allele. A change of amino acid 293 from aspartic acid to asparagine renders the protein non-functional, as demonstrated by expression in S. cerevisiae zrc1cot1 mutant cells (Figure S3). D293 is highly conserved in cation diffusion facilitator (CDF) proteins from bacteria to humans and was recently found to be essential for Zn2+ transport activity when expressed in zrc1cot1 (Kawachi et al., 2012). Our data provide in planta support for this finding. Furthermore, the strength of the ozs1 Zn-hypersensitivity phenotype confirms the major role of AtMTP1 in Zn2+ tolerance at the seedling stage (Kobae et al., 2004; Desbrosses-Fonrouge et al., 2005).

The ozs2 mutant carries a mutation in AtPME3

Specific Zn2+ hypersensitivity was also found for ozs2. In contrast to ozs1 as well as the mutants ozs3-6, the mutation in ozs2 is semi-dominant. It was mapped to an amino acid change at position 497 (glycine to valine) in AtPME3. While in accordance with the semi-dominant character no complementation with wild-type AtPME3 was achieved and two T-DNA insertion lines showed wild-type Zn2+ tolerance, the available evidence strongly suggests that the change is causal for the ozs2 phenotype. First, no other mutation was found in the mapped interval. Second, and most importantly, down-regulation of PME3 transcript levels by RNAi fully restored wild-type Zn2+ tolerance in ozs2. Third, overexpression of the mutated PME3 version in wild-type plants led to severe and specific Zn2+ hypersensitivity. Surprisingly, overexpression of the Col-0 version was equally detrimental (Figure 6a). For both AtPME3 versions we found a remarkably strong correlation between transcript abundance and degree of growth inhibition by excess Zn2+ (Figure 6c,d).

Plant PMEs are encoded by large gene families. The A. thaliana genome contains 66 PME genes, the poplar genome 89 (Pelloux et al., 2007). Pectin methylesterases catalyze the demethylesterification of homogalacturonan pectin subsequent to its secretion into the cell wall in a highly methylesterified state. This process plays a crucial role in the regulation of cell elongation as it greatly influences cell wall architecture and extensibility (Peaucelle et al., 2011a). Accordingly, important functions in vegetative and reproductive development have in recent years been assigned to the regulated modulation of methylesterification of homogalacturonan by PMEs (Wolf et al., 2012). For instance, the tip growth of pollen tubes is dependent on spatially controlled PME activity (Bosch and Hepler, 2005). Regulated pectin demethylesterification is an early and necessary event in phyllotaxis and organ initiation (Peaucelle et al., 2008, 2011a). In addition, PMEs have been studied in the context of plant–pathogen interactions (Lionetti et al., 2012). An increase in the number of free carboxylic groups renders pectin more susceptible to degradation through polygalacturonases secreted by pathogens.

However, few individual PMEs have been functionally characterized to date. Most mutants do not show phenotypes, probably because of overlapping activities of PMEs as well as compensatory responses (Wolf and Greiner, 2012). Phyllotaxis requires the differential regulation of AtPME5 in the shoot apical meristem and the elongating stem by the transcription factor BELLRINGER (Peaucelle et al., 2011b). Recently, a contribution of AtPME35 to mechanical strength of the stem was demonstrated (Hongo et al., 2012). AtPME3 was identified as a protein interacting with and possibly targeted by a cellulose-binding protein from the parasitic nematode Heterodera schachtii (Hewezi et al., 2008). Later, a role of AtPME3 in controlling adventitious root formation was described (Guénin et al., 2011).

AtPME3 is strongly expressed throughout the plant (Louvet et al., 2006). According to publicly available microarray data transcript levels are highest in the vascular tissue of roots and shoots (http://bar.utoronto.ca/efp/cgi-bin/efpWeb.cgi). This is consistent with promoter::GUS data (Guénin et al., 2011). Mature AtPME3 protein, i.e. without the pro-region, has been detected in the cell wall (Boudart et al., 2005; Guénin et al., 2011).

Two major questions arise from the observations reported for the ozs2 mutant: (i) what is the effect of the mutation on the biological activity of AtPME3; and (ii) why does the ozs2 mutation as well as the ectopic expression of AtPME3 cause severe and specific Zn2+ hypersensitivity? The ozs2 mutation changes a highly conserved stretch of amino acids. The mutated glycine is nearly 100% conserved in PMEs across kingdoms (Markovic and Janecek, 2004). It is located next to an arginine that is part of the active center (Johansson et al., 2002), suggesting a change in enzyme activity of ozs2 AtPME3. It was not possible to directly test this hypothesis because no purified recombinant protein representing the PME domain could be obtained, a common problem with type-I PMEs (De-la-Peña et al., 2008). Instead, we tested whether the deleterious effect of ozs2 AtPME3 is dependent on enzyme activity by mutating two residues that, according to structural mechanistic knowledge and experimental validation (Dorokhov et al., 2006), lead to an enzymatically dead protein. For overexpressing plants we found at least equally pronounced growth inhibition by excess Zn2+ as for the ozs2 mutant version (Figure 6b), arguing strongly against an effect on activity as the underlying cause of the ozs2 phenotype. Consistent with this hypothesis, we did not detect any differences in total PME activity between any of the studied plant lines, regardless of pH, assay type or substrate.

A detectable biochemical consequence of the ozs2 mutation, however, concerned the proteolytic processing that type-I PMEs undergo before they reach the apoplast. The fraction of mature protein was consistently smaller for the ozs2 version compared with the wild-type AtPME3 (Figure 7). This difference was independent of Zn2+ exposure. While the ozs2 mutation is not directly located in one of the two basic cleavage motifs of the pro-protein (Wolf et al., 2009), the results still suggest that proteolytic processing is impaired in the mutant. It is widely assumed that PMEs are processed intracellularly, most likely before exit from the Golgi (Wolf et al., 2009). Possibly the processing of PMEs occurs in Golgi-localized protein complexes similar to those postulated recently for cell wall biosynthesis enzymes (Burton et al., 2010; Oikawa et al., 2013). We speculate that the semi-dominant ozs2 mutation affects the interaction of AtPME3 with proteases or other proteins along the secretory pathway. Accordingly, we hypothesize that strong overexpression of AtPME3 interferes with the function of proteins trafficking through the Golgi in a similar fashion. Both impaired processing and protein overdose would then indirectly affect synthesis and/or modification of the cell wall in a way that renders seedlings hypersensitive to Zn2+.

Two observations indicated alterations in the cell walls of ozs2 seedlings. First, upon exposure to Zn2+ ozs2 showed more extensive root hair outgrowth (Figure 3). This is consistent with the role of the pectin network in controlling cell expansion. Cell wall extensibility is the key parameter, and pectins as the most complex matrix components play a major role in the modulation of the physical properties of the cell wall (Wolf and Greiner, 2012). Accumulation of Zn in roots, however, was not affected by the change in root hair growth, arguing against an effect on Zn uptake.

Second, an excess of Ca2+ was able to suppress the Zn2+ hypersensitivity of ozs2 but not of ozs1 seedlings (Figure 8). This finding pointed towards alterations in cation-binding sites in the apoplast of ozs2 as the cause for the phenotype. Free carboxylic groups of demethylesterified pectin could potentially bind metal cations and thereby lower the uptake into the symplast. As an example, 30% of the total Zn in roots of the metal hyperaccumulator Noccaea caerulescens was estimated to be localized in cell walls (Salt et al., 1999). Also, it is documented in vitro that pectin with lower degree of methylation has a higher Zn-binding capacity (Khotimchenko et al., 2008). Increases in low-methylesterified pectin have been implicated in metal tolerance (Krzeslowska, 2011). Thus, it is suggestive to associate a loss of metal tolerance with a defect in demethylesterification of pectin. However, the specificity of the metal hypersensitivity of ozs2 strongly argues against a simple change in the metal-binding capacity of the cell wall. Other metal cations such as Cd2+ or Cu2+ should also bind to pectins and therefore potentially affect growth of ozs2. In fact, Cu2+ is known to bind more strongly to pectins than Zn2+. Accordingly, in a recent synchrotron-based x-ray fluorescence microscopy/x-ray absorption spectroscopy study with metal-exposed roots of cowpea, Cu was–in contrast to Zn–found to be mainly associated with polygalacturonic acids (Kopittke et al., 2011).

Moreover, careful analysis of Col-0 and ozs2 roots via immunostaining of pectins and electron microscopy did not reveal any consistently noticeable differences between the cell walls of the two genotypes, regardless of Zn2+ status. Taken together, our results do not agree with a modification of cell wall-binding strength as the reason for the ozs2 Zn2+ hypersensitivity.

Instead, we hypothesize that due to subtle changes caused by the ozs2 mutation the interference of Zn2+ ions with the complex dynamics of cell wall architecture is aggravated. Our observations suggesting changes in cell wall properties of ozs2 were restricted to seedlings exposed to Zn2+, indicating that Zn2+ can act as a trigger for cell wall remodeling. This adds to recent evidence for the effects of Zn on root development and morphology that could be due to effects on cell wall modulation. An analysis of natural variation in Zn tolerance in A. thaliana found indications that Zn is required for the initiation of lateral roots (Richard et al., 2011). Some A. thaliana accessions did not produce lateral roots under conditions of Zn2+ deficiency. A study on responses to Zn2+ excess on the other hand found effects on the shape of A. thaliana root hairs (Fukao et al., 2011). The ozs2 mutant might therefore help identify primary Zn2+-toxicity targets which are currently unknown (Clemens, 2010).

Furthermore, the ozs2 mutation provides opportunities to better understand cell wall architecture and remodeling. The cell wall in general is biochemically poorly understood. Up to 10% of all A. thaliana genes are estimated to be involved in cell wall biosynthesis, modification, degradation, and the regulation of these processes (Liepman et al., 2010). Structural analysis of pectins as the most complex polysaccharides in cell walls is particularly challenging, and pectin biosynthesis is the least understood (Mohnen, 2008). Complexity is potentiated through the existence of a large number of modifying enzymes including the PMEs. Their activity may promote a multitude of apparently contradictory changes including wall stiffening, wall loosening, wall degradation and wall signaling (Wolf et al., 2012). An important aspect of PME regulation is the proteolytic cleavage of the pro-domain (Wolf et al., 2009). The processing defect in the ozs2 AtPME3 can be used to unravel this aspect of control of PME activity. Moreover, the amazingly tight correlation between AtPME3 overexpression levels and the degree of Zn2+ hypersensitivity (Figure 6c,d) could be exploited to trigger and analyze changes in cell wall structure varying in magnitude over a wide range.

Experimental Procedures

Plant material

All A. thaliana plants analyzed in this study were in the Col-0 background. The EMS-mutagenized M2 seeds were purchased from Lehle Seeds (http://www.arabidopsis.com/). The atpme3 T-DNA insertion line (Gabi-Kat 002A10) was obtained from the Gabi-Kat collection (University of Bielefeld, Germany). Wild-type and ozs2 mutant AtPME3 coding sequences were cloned into pMDC83 (Curtis and Grossniklaus, 2003) via the Gateway system (Invitrogen, http://www.invitrogen.com/). The 396-bp DNA-fragment used for RNAi was amplified from cDNA, cloned into pENTR/D-Topo (Invitrogen) and subcloned into pHELLSGATE8 (Helliwell et al., 2002). For expressing the C-terminally HA-tagged versions the coding sequence was amplified from cDNA (for primers see Table S2), ligated into pSGP72 and then with the HA tag into pMDC32.

For complementation of ozs1 a genomic fragment consisting of promoter, coding sequence, and terminator was PCR amplified (primers see Table S2) and subcloned into pCB302.

All plasmids used for the generation of transgenic A. thaliana lines were transformed into Agrobacterium tumefaciens strain GV3101. Transformation of A. thaliana was performed via the floral dip method.

Plant cultivation and tolerance assays

For vertical plate tolerance assays surface-sterilized seeds were placed on agar plates containing modified 1/10 Hoagland's medium [0.28 mm Ca(NO3)2, 0.1 mm (NH4)H2PO4, 0.2 mm MgSO4, 0.6 mm KNO3, 5 μm of a complex of Fe(III) and N,N′-di-(2-hydroxybenzoyl)-ethylenediamine-N,N′-diacetate (HBED; ABCR GmbH, http://www.abcr.de/), 5 mm MES, 1% (w/v) sucrose, 1% (w/v) Type-A agar (Sigma-Aldrich, http://www.sigmaaldrich.com/), pH 5.7]. Different metal salts, paraquat, mannitol, or NaCl were added to the medium. After about 48 h at 4°C (in the dark) plates were transferred to 23°C and cultivated under long-day conditions (16 h light/8 h dark). The length of the primary root of each seedling was measured after about 12 days (when seedlings on control plates had reached the bottom of the plate). Seedlings were also cultivated in liquid modified 1/10 Hoagland's medium in Falcon tubes at 80 r.p.m.

Elemental analysis

Shoots and roots were harvested separately and rinsed four times (Millipore water, 2 × 20 mm CaCl2, Millipore water) for 10 min at 4°C. After lyophilization plant material was digested in a 2:1 mixture of HNO3 (65%, v/v) and H2O2 (30%, v/v) in a microwave oven using a temperature step gradient (maximum of 210°C). Samples were analyzed by inductively coupled plasma optical emission spectrometry on an iCAP 6500 Series spectrometer (Thermo-Fisher, http://www.thermofisher.com/).

Transcript analysis

The RNA was extracted from homogenized plant material with TRIzol (Invitrogen) according to the manufacturer's instructions. The quality of the RNA was checked with a NanoPhotometer (IMPLEN, http://www.implen.de/). DNaseI-treated RNA (1.0 μg) was used for cDNA synthesis (RevertAid First Strand cDNA Synthesis Kit, Thermo-Fisher). Quantitative real-time PCR (qRT-PCR) reactions were performed in 96-well plates in a Bio-Rad iCycler with a MyiQ real-time PCR detection system using SYBR Green (iQ SYBR Green supermix, Bio-Rad, http://www.bio-rad.com/) to monitor cDNA amplification. Five microliters of 1:50 diluted cDNA and 5 pmol of forward and reverse primers were added to 10 μl SYBR Green mix in a total volume of 20 μl. The standard thermal profile was 95°C for 10 min, followed by 40 cycles of 95°C for 10 sec and 60°C for 1 min. Data were analyzed using iQ5 Optical System software version 2.1 (Bio-Rad). The ΔCT value was calculated by subtracting the cycle threshold (CT) value of the reference gene EF1α from the CT value of the target gene PME3. The relative transcript level (RTL) was calculated as follows: RTL = 1000 × 2−ΔCT. Primers (Table S2) were designed using the Primer3 software (http://primer3.sourceforge.net/). Primers and amplicons were checked for potential secondary structures with the program mfold (http://mfold.rna.albany.edu/?q=mfold).

Protein detection

Protein was extracted according to Wolf et al. (2009). Following SDS-PAGE, proteins were transferred to a nitrocellulose membrane. Primary antibody (anti-HA, H9658, Sigma-Aldrich) was diluted 1:3000 and secondary antibody (anti-mouse, A9044, Sigma-Aldrich) coupled to peroxidase was diluted 1:10 000 with 5% (w/v) milk powder in 2-amino-2-(hydroxymethyl)-1,3-propanediol (TRIS)-buffered saline and Tween 20. Signals were detected using an ECL kit (GE Healthcare, http://www.gehealthcare.com/). For quantification of signal intensities ImageJ software (V 1.43u, Rasband, 1997–2006) was used.

Yeast transformation and growth assay

For growth assays the Zn2+-hypersensitive S. cerevisiae strain zrc1 cot1 was used. Wild-type and ozs1 mutant AtMTP1 coding sequences were first cloned into pSGP72 to add a C-terminal HA tag and from there into the vector pYES2. Saccharomyces cerevisiae cells were grown at 30°C in yeast nitrogen base supplemented with the appropriate amino acids and carbohydrates. Tolerance assays in liquid culture were performed as described in Tennstedt et al. (2009).

Pectin methylesterase activity assays

Plants were grown using the hydroponic cultivation system. Roots were harvested and ground in liquid nitrogen. Total soluble protein was extracted according to Guénin et al. (2011). Ten micrograms of soluble protein was used for activity assay. The PME activity was measured with the methanol oxidation assay according to Reca et al. (2012). The assay was performed at a pH of 7.5 and with citrus pectin (P9135, Sigma-Aldrich) as the substrate. A possible influence of the degree of pectin methylation was assessed by also testing highly methylesterified pectin (>85%, P9561, Sigma-Aldrich) in parallel. The reaction was monitored for 60 min with a microplate reader (PowerWave × 340, BioTek, http://www.biotek.com/).

Alternatively, PME activity was determined using the pH indicator assay (Hewezi et al., 2008) and the gel diffusion assay (Downie et al., 1998). For both assays citrus pectin (P9135, Sigma-Aldrich) was also used as the substrate. The pH indicator assay was performed at a pH value of 7.0. For the gel diffusion assay the substrate was dissolved in 0.1 m citrate/0.2 m phosphate (dibasic) buffers adjusted to pH 5.0, 6.0, 7.0, and 8.0. After addition of 1% agar and autoclaving, Petri dishes were prepared: 4 μg of soluble protein in 20 μl was added to wells punched in the gel. As a negative control protein extracts were boiled for 5 min and used in parallel.

Confocal microscopy

Five-day-old roots were incubated in a propidium iodide (10 mg l−1) solution to stain the cell wall. Propidium iodide fluorescence was visualized with a Leica SP5 confocal laser-scanning microscope (excitation 561 nm, emission 580–650 nm; http://www.leica-microsystems.com/). Cell size and cell numbers were determined using ImageJ-software.

Electron microscopy

Seedling roots were cut into segments of length 2 mm and fixed in 100 mm Na-cacodylate containing 2.5% glutaraldehyde and 3% formaldehyde, pH 7.3, for 4 h at 4°C and further processed as described in Breuers et al. (2012).


We are grateful to Christiane Meinen, Pia Schuster, and Doris Wittmann for excellent technical assistance, and to Christina Reitmaier-Weber for scientific discussion. We thank the Genetics Department at the University of Bayreuth for access to the confocal microscope.