*Article* **Isolation and Characterization of Copper- and Zinc-Binding Metallothioneins from the Marine Alga** *Ulva compressa* **(Chlorophyta)**

**Antonio Zúñiga 1,2, Daniel Laporte 1,\*, Alberto González 1, Melissa Gómez 1, Claudio A. Sáez 2,3 and Alejandra Moenne 1,\***


Received: 23 September 2019; Accepted: 22 December 2019; Published: 25 December 2019

**Abstract:** In this work, transcripts encoding three metallothioneins from *Ulva compressa* (UcMTs) were amplified: The 5 and 3 UTRs by RACE-PCR, and the open reading frames (ORFs) by PCR. Transcripts encoding UcMT1.1 (*Crassostrea*-like), UcMT2 (*Mytilus*-like), and UcMT3 (*Dreissena*-like) showed a 5 UTR of 61, 71, and 65 nucleotides and a 3 UTR of 418, 235, and 193 nucleotides, respectively. UcMT1.1 ORF encodes a protein of 81 amino acids (MW 8.2 KDa) with 25 cysteines (29.4%), arranged as three motifs CC and nine motifs CXC; UcMT2 ORF encode a protein of 90 amino acids (9.05 kDa) with 27 cysteines (30%), arranged as three motifs CC, nine motifs CXC, and one motif CXXC; UcMT3 encode a protein of 139 amino acids (13.4 kDa) with 34 cysteines (24%), arranged as seven motifs CC and seven motifs CXC. UcMT1 and UcMT2 were more similar among each other, showing 60% similarity in amino acids; UcMT3 showed only 31% similarity with UcMT1 and UcMT2. In addition, UcMTs displayed structural similarity with MTs of marine invertebrates MTs and the terrestrial invertebrate *Caenorhabtidis elegans* MTs, but not with MTs from red or brown macroalgae. The ORFs fused with GST were expressed in bacteria allowing copper accumulation, mainly in MT1 and MT2, and zinc, in the case of the three MTs. Thus, the three MTs allowed copper and zinc accumulation in vivo. UcMTs may play a role in copper and zinc accumulation in *U. compressa*.

**Keywords:** copper and zinc; expression in bacteria; metal accumulation; metallothioneins; marine alga; *Ulva compressa*

### **1. Introduction**

Metallothioneins (MTs) are low molecular weight proteins, of around 10 kDa, that are rich in cysteine residues allowing the binding of divalent or monovalent metal ions such as Zn2<sup>+</sup>, Cd2<sup>+</sup>, Pb2<sup>+</sup>, Hg2+, Cu1+, Ag1+, among others [1–3]. MTs participate in metal accumulation and detoxification in vertebrates, invertebrates, plants, algae, and bacteria [1–3]. Cysteine residues in MTs are usually arranged as CC, CXC, and/or CXXC motifs and they correspond to around 30% of amino acids. In vertebrates, cysteine residues in MTs are contained in two domains, α and β, that are separated by a linker of variable sizes. Vertebrate MTs are rich in glycine and alanine amino acids, ranging from 10% to 20% residues [2] and invertebrate and plant MTs can contain histidine and aromatic residues [4,5]. MTs were first discovered in horse kidney, although they have been isolated from kidney and liver

of other mammals [6]. In mammals, such as humans and mouse, there are four MT isoforms and the linker is constituted by three amino acids [2]. In fish, such as in rainbow trout, there are two MTs and the linker is formed by four amino acids [7]. In invertebrates, there are mainly two MTs, and the cysteine-rich domains are separated by a linker of four to seven amino acids and they have been described in organisms such as the nematode *Caenorhabtidis elegans* [8], the gastropod *Arianta arbustorum* [9], the snail *Helix pomatia* [10,11], the mollusks *Crassostrea virginica* [12], *Mytilus edulis* [13], and *Dreissena polymorpha* [14] and in the equinodern *Strongylocentrotus purpuratus* [15]. The yeast *Saccharomyces cerevisiae* displays two MTs, CUP-1 and CRS5, and the linker is constituted by three amino acids [16,17]. The cyanobacteria *Synechococcus* sp. showed a single MT, SmtA, and the linker that separates the two cysteine-rich domains is constituted by 15 amino acids [2]. The first MT isolated in plants was wheat germ MT, the first cloned MT in plants was *Mimulus guttatus MT*, and the first cloned MT from marine algae was *Fucus vesiculosus* MT [18–20]. Plant MTs can be classified in four types corresponding to type 1, 2, 3, and 4 [3]. The types 1, 2, and 3 MTs have cysteine-rich domains separated by a linker of around 40 amino acids and a linker of about 15 amino acids, and they differ in the arrangement of cysteines in cysteine-rich domains [3]. Type 4 MTs have mainly a linker of around 15 amino acids [3]. In *Arabidopsis thaliana,* there are seven MTs corresponding to MT1a, MT1c, MT2a, MT2b, MT3, MT4a, and MT4b and in *Populus trichcarpa x deltoides* there are six MTs corresponding to MT1a, MT1b, MT1a, MT1b, MT3a, and MT3b [21,22]. In addition, the study of algal genomes has shown that the brown macroalgae (Phaeophyceae) *Fucus serratus*, *Ectocarpus siliculosus*, and *Sargassum binderi*, and the red macroalgae (Rodophyceae) *Chondrus crispus* and *Euchema denticulatum* encode a single MT [3]. Red and brown algae MTs are related among each other considering the arrangement of cysteines [3]. Until now, no MTs have been cloned or characterized in green macroalgae (Chlorophyceae).

The green macroalga *Ulva compressa* is the dominant species in copper-polluted coastal areas of northern Chile and in other parts of the world [23,24]. It has been shown that the alga collected in the field accumulate copper in its tissue [23]. Furthermore, the alga cultivated in vitro with 2.5, 5, 7.5, and 10 μM for 0 to 12 days showed a linear accumulation of intracellular copper with increasing concentrations of the metal reaching a maximal accumulation of 620 μg g−<sup>1</sup> of dry weight (DW), at day 12 with 10 μM copper [25]. In addition, *U. compressa* extrudes copper ions to the extracellular medium reaching a maximal concentration at intracellular level of around 900 μg g−<sup>1</sup> of DW [24]. In contrast, the green alga *Ulva fasciata* cultivated with 0.3 μM copper for 14 days reached an intracellular level of copper of 2000 μg g−<sup>1</sup> of DW suggesting that this alga may not extrude copper ions to the culture medium [26]. On the other hand, the level of MT transcripts in *U. compressa* cultivated with 7.5 and 10 μM of copper increased from days 3 to 12 [24]. Thus, it is possible that accumulation of intracellular copper is mediated by MTs in *U. compressa*. In this sense, it has been shown that *Arabidopsis thaliana* plants deficient in MT1a accumulate 30% less copper in the shoots than control plants [26] and *A. thaliana* mutants deleted in four MTs accumulate 45% less copper in the shoots [27]. In addition, rat fibroblasts having a deletion of MTI and MTII accumulate less copper than control cells and is subjected to an increase in oxidative stress [28]. Thus, it is possible that *U. compressa* can accumulate copper mediated by MTs. A transcriptomic analysis performed with the alga cultivated with 10 μM copper for 0 and 24 h allowed the identification of 7 potential MTs in *U. compressa* and their levels increased from days 3 to 12 of metal exposure [29].

In this work, we cloned three putative *U. compressa* MTs previously described as *Crassostrea*-like, *Mytilus*-like, and *Dreissena*-like MTs [26], which were renamed UcMT1, UcMT2, and UcMT3, respectively. In this work, the ORF of each UcMT was cloned and expressed in the bacteria *E. coli* as a fusion protein with a glutahione-S-transferase (GST) allowing accumulation of copper and zinc in vivo. Thus, the marine alga *U. compressa* may accumulate intracellular copper and zinc through UcMTs.

### **2. Results**

### *2.1. Sequences of Transcripts Encoding UcMTs*

Total RNA and mRNAs were isolated from *U. compressa* cultivated with 10 μM copper for three days. The 5 and 3 untranslated regions (UTR) of transcripts encoding three UcMTs were amplified using RACE-PCR technique, as well as the open reading frame (ORF) using conventional PCR. UcMT1.1 transcript (formerly *Crassostrea*-like *mt*) and protein are described in Table 1 (Figure 1). It is important to mention that two other transcripts that were closely related with UcMT1.1 were isolated, corresponding to UcMT1.2 and UcMT1.3; UcMT1.2 showed a deletion of 48 nucleotides after G in position 471 of the 3´UTR region of UcMT1.1, and UcMT1.3 displayed the same deletion mentioned before and a deletion of 58 nucleotides after C in position 607 in the 3´UTR of UcMT1.1 (Figure 1). UcMT2 transcript (formerly *Mytilus*-like *mt*) and protein is described in Table 1 (Figure 2) as well as UcMT3 transcript (formerly *Dreissena*-like *mt*) and protein (Figure 3). The linker region of UcMT1.1, UcMT2, and UcMT2 correspond to, 9, 9, and 23 amino acids, respectively (Figures 1–3).


**Figure 1.** Complete cDNA and amino acid sequences of metallothionein UcMT1.1 (*Crassostrea*-like) from *U. compressa*. Initiation and termination codons are highlighted in bold; \* indicate stop codon, cysteines arranged as CXC and CC motifs are underlined in the amino acid sequence of UcMT1. UcMT1.2 presents a deletion of 48 nucleotides located after the G in position 471 in the 3 UTR of MT1.1, and the deletion is underlined. UcMT3 present the deletion previously mentioned, and a deletion located after the C in position 607 in the 3 UTR of UcMT1.1, and the deletion is underlined.


**Table 1.** Characteristics of *U. compressa* metallothioneins (UcMTs) transcripts and proteins. For transcripts: Nucleotides (nt); for proteins amino acids (aa) and cysteines (cys).

**Figure 2.** Complete cDNA and amino acid sequences of metallothionein UcMT2 (*Mytilus*-like) from the marine alga *U. compressa*. Initiation and termination codons are highlighted in bold; \* indicate stop codon, cysteines arranged as CXC, CC, and CXXC motifs are underlined in the amino acid sequence of UcMT2.


**Figure 3.** Complete cDNA and amino acid sequences of metallothionein UcMT3 (*Dreissena*-like) from the marine alga *U. compressa*. Initiation and termination codons are highlighted in bold; \* indicate stop codon, cysteines arranged as CXC and CC motifs are underlined in the amino acid sequence of UcMT3.

### *2.2. Similarities in Amino Acids of UcMTs and Hierarchical Clustering of MTs*

UcMT1.1 and UcMT2 were more closely related among each other and shared 52.2% identity and 60% similarity in amino acids; indeed, both sequences contained arrangement of cysteines corresponding to 3 CC and 9 CXC motifs, but UcMT2 also contained a CXXC motif (Figure 4A). In contrast, UcMT3 shared only 30% similarity in amino acids with MT1.1 and MT2 and contained arrangements of cysteines corresponding to 7 CC and 7 CXC (Figure 4A). UcMT3 showed an extra N-terminal sequence of 10 amino acids, an internal additional sequence of 19 amino acids, and an extra C-terminal sequence of 8 amino acids, compared with UcMT1.1 and UcMT2 (Figure 4A). Thus, UcMT1 and UcMT2 may have derived from UcMT3 by deletions of initial, internal, and terminal nucleotide sequences.

The hierarchical clustering of vertebrate, invertebrate, and plants MTs constructed with 237 protein sequences (including the 3 UcMTs) demonstrated that UcMT1.1 and UcMT2 grouped mainly with marine crustacean MTs, such as those of the lobster *Homarus americanus* and the crabs *Carcinus maena* and *Scylla serrata* (Figure S1). In addition, UcMT3 clustered with the nematode *C. elegans* MTs, as well as with MTs of marine equinoderms MTs such as those of *Sterechinus neumayeri*, *Strongylocentrus purpuratus,* and *Paracentrotus livudus* (Figure S1). On the other hand, UcMTs grouped as a different clade with MTs from Rodophyceae and Phaeophyceae (Figure 4B). Thus, *U. compressa* MTs are more

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closely related with marine invertebrate MTs and the terrestrial invertebrate *C. elegans* MTs, and not to MTs from other marine macroalgae.

**Figure 4.** Alignment of amino acid sequences of metallotioneins (MTs) UcMT1.1, UcMT2, and UcMT3 from the marine alga *U. compressa* (**A**). Identical amino acids are indicated in black and similar amino acids are indicated in gray. \* indicates identical amino acids and · similar amino acids. Hierarchical clustering of the amino acid sequences of UcMTs and MTs found in other marine macroalgae (**B**).

### *2.3. Expression of UcMTs-GST in Bacteria and Detection of GST-Tag*

The ORFs of UcMTs were cloned in an *E. coli* expression vector, which allows the expression of MTs fused with the enzyme glutathione-S-transferase (GST) from the platyhelminthe *Schistosoma japonicum* (26 kDa), an enzyme that contain a single cysteine in the N-terminal domain, and do not bind metals. After 1 to 12 h of culture, the induction of protein expression with IPTG allows the visualization of increasing levels of UcMT1.1-GST (34.2 kDa. Figure 5A), UcMT2-GST (35.05 KDa, Figure 5B), and UcMT3-GST (39.4 KDa, Figure 5C). These proteins were detected by Western blot using an antibody prepared against *S. japonicum* GST indicating that the overexpressed proteins correspond to UcMTs fused with GST (Figure 5C).

**Figure 5.** Visualization of protein extracts of bacteria overexpressing UcMT1.1-GST (**A**), UcMT2-GST (**B**), and UcMT3-GST (**C**) obtained from bacteria cultivated without IPTG (lane 1) or with 0.5 mM IPTG for 1, 3, 6, 9, and 12 h (lanes 2 to 7). Visualization of proteins bacterial extracts overexpressing UcMTs-GST transferred to a nitrocellulose membrane stained with Ponceau red dye (**D**) or incubated with anti-GST antibody and revealed by chemiluminescence (**E**). Proteins of extracts overexpressing GST (lane 1), not overexpressing GST (lane 2), and overexpressing UcMT1.1-GST (lane 3), UcMT2-GST (lane 4), and UcMT3-GST (lane 5) for 6 h. Arrows (left side) indicate molecular weights of standard proteins and arrows (right side) indicate molecular weights of UcMT3-GST (39.4 kDa), UcMT2-GST (35.05 kDa), UcMT1 (34.2 kDa), and GST (26 kDa).

### *2.4. UcMTs-GST-Mediated Accumulation of Copper or Zinc In Vivo*

Transformed bacteria expressing MTs-GST were cultivated with 0.5 mM IPTG for 30 min, and with 1 mM copper and IPTG for 6 h. Control bacteria expressing only GST accumulate 0.35 μg of copper mg−<sup>1</sup> of dry weight (DW), whereas those expressing UcMT1.1-GST, UcMT2-GST, and UcMT3-GST accumulate 1.8, 1.7, and 1.4 times more copper than the control, respectively (Figure 6A). On the other hand, transformed bacteria were cultivated with 0.5 mM IPTG for 30 min and, with 1 mM zinc and IPTG for 6 h. Control bacteria accumulate 0.49 μg mg−<sup>1</sup> of zinc mg−<sup>1</sup> of DW whereas those expressing UcMT1.1-GST, UcMT2-GST, and UcMT3-GST accumulate 4,1, 3.8, and 3.4 times more zinc than the control, respectively; although these increases were not significantly different among each other (Figure 6A). Thus, expression of UcMTs-GST mediates accumulation of copper and zinc in vivo.

**Figure 6.** Level of copper (**A**) and zinc (**B**) in *E. coli* cultivated without IPTG for 6 h (control) and with 0.5 mM IPTG for 6 h and overexpressing UcMT1.1, UcMT2, and UcMT3. Levels of copper and zinc are expressed in micrograms per gram of bacterial dry weight (DW). Bars represent mean values of three independent experiments ± SD. Letters indicate significant differences among mean values (*p* < 0.05).

### **3. Discussion**

In this work, we isolated the complete sequence of transcripts encoding three MTs from the marine alga *U. compressa*. These transcripts encode UcMT1.1, UcMT2, and UcMT3 with a MW of 8.3, 9.05, and 13.4 kDa, respectively, and containing 24–30% of cysteines. It is important to mention that UcMT1.1 and UcMT2 seemed to be structurally more related among each other than to UcMT3. UcMT1.1 and UcMT2 showed deletions in amino acids sequences compared with MT3 at the initial, internal, and terminal part of the protein compared with UcMT3. Thus, UcMT2 and UcMT3 genes may have derived from UcMT3 gene. In the case of UcMT1, it seems that more than a single copy exists in *U. compressa* genome since three different transcripts of UcMT1 were isolated corresponding to UcMT1.1, UcMT1.2 and UcMT1.3. In this sense, it has been shown that the increase in number of copies of Cup1 MT gene in the yeast *S. cereviciae* allowed an increased tolerance to copper [30]. In addition, several copies of domains α and β of MTs exist in the mollusk *C. gigas* [12]. Thus, it is not surprising that UcMT1 may be present in multiple copies in *U. compressa* genome, but the latter remained to be confirmed. The sequencing of *U. compressa* genome is already in course.

Interestingly, UcMTs resemble marine invertebrate MTs. Mollusk MTs are constituted by 73–75 amino acids and contain 20–21 cysteine residues (28%), arranged mainly as CXC and CC motifs [5]. UcMTs are longer than mollusk MTs since they are constituted by 81, 90, and 139 amino acids; although the content of cysteines is similar to mollusk MTs (28–30%). However, mollusk MTs reported until now do not contain histidine or aromatic residues [5]. In addition, MT from marine crustacean and equinoderms showed cysteines (29–30%) arranged as CC and CXC and they do not contain histidine or aromatic aminoacids [25,26]. It is important to mention that we showed that UcMT1.1 contains a tyrosine, UcMT2 a histidine, and UcMT3 a tryptophan. In this sense, the two MTs found in the worm *C. elegans* are constituted by 75 amino acids and contain 19 cysteines (25%), arranged as CXC and CC, and contain a tyrosine and histidine residues [8]. Thus, UcMTs are more similar to *C. elegans* MTs but longer and contain a higher percentage of cysteines (28–30%). This indicates that UcMTs are unique MTs, and their sequences are more closely related with MTs of marine invertebrates and *C. elegans* MTs. It is not surprising that UcMTs showed similarities to marine invertebrate MTs since *F. vesiculosus* MT is also more related to mollusk MTs than to plant MTs [20]. *F. vesiculosus* MT showed a linker of 14 amino acids, which is longer than the linker of vertebrates MTs (4 aa) and the linkers of plant MTs (4 and 40 aa) [20]. The linker of UcMTs has a size of 9–13 amino acids that is more closely related with *F. vesiculosus* and invertebrate MTs linker size.

In addition, the sequence of UcMTs is different from the MT cloned from *F. vesiculosus*, which is constituted by 67 amino acids (6.9 kDa) and contains 16 cysteine residues (24%) arranged as CXC, but not as CC [3]. The only structural similarity of *Fucus mt* is that this transcript showed a short 5 UTR of 64 nucleotides and a long 3 UTR of 960 nucleotides [8]. Furthermore, MTs identified in the genomes of Rodophyceae *C. cripus* and *E. denticulatum* are constituted by 69 to 71 amino acids and contain 12 to 14 cysteines (20%) arranged mainly as CXC, but not as CC [2]. Furthermore, UcMTs grouped in a different clade compared with MTs from other marine macroalgae. Thus, UcMTs of the green macroalga *U. compressa* are distantly related with those found in red and brown macroalgae.

On the other hand, we showed that UcMTs-GST mediates the accumulation of copper and zinc in bacteria. In particular, UcMT1.1 and UcMT2, which are structurally more closely related, allowed higher accumulation of copper compared with UcMT3. Moreover, the three MTs allowed accumulation of zinc with similar efficiencies among them. Thus, the three UcMTs differentially bind copper and similarly bind zinc in vivo but their affinity for copper and zinc need to be further investigated. In this sense, it has been demonstrated that MTs are either more Zn- or Cu-binding thioneins, preference associated with cysteine arrangements and the nature of the other amino acids that constitute the MT [4,31]. Likewise, mouse MT1, MT2, and MT3 are more Zn-thionein and, in contrast, yeast cup-1, *Drosophila* MntA and MntB, and mouse MT4, are more Cu-thioneins [4,31]. Thus, the nature of UcMTs regarding their affinity for copper or zinc need to be further investigated.

It is now clearly established that green and red macroalgae are more closely related among each other, and with terrestrial plants, than with brown macroalgae [32–34]. Green and red macroalgae belong to the kingdom Plantae, as terrestrial plants [35] whereas brown macroalgae belong to the kingdom Chromalveolta [34]. The latter is based on the observations that green and red algae contain a plastid that derive from a single event of endosymbiosis by a cyanobacteria, whereas brown algae plastids derive from a secondary or tertiary endosymbiosis event of green or red microalgae [32,33]. Thus, it is unexpected that the three UcMTs of the green macroalga *U. compressa* are not closely related with other marine alga MTs, in particular to red macroalgae MTs. In contrast, the major similarity is with MTs of marine invertebrate and *C. elegans* MTs. Considering that marine algae appeared on earth around a billion years ago, and marine invertebrates emerged around 500 million years ago [34,36] and, moreover, that UcMTs are longer than MTs of marine invertebrates, it is then possible that marine invertebrates have acquired MTs genes from marine green macroalgae by horizontal gene transfer; however, this hypothesis need to be further investigated. Furthermore, it is possible that *U. compressa* contain additional MTs that differentially bind copper, zinc, and other heavy metals, as it has been predicted in [23], but the latter need to be further analyzed. In this sense, it has been shown that the equinoderm *Paracentrotus livudus* exhibits 7 MTs [35] suggesting that the four other potential UcMTs can be functional MTs. Thus, additional UcMTs may exist in the genome of *U. compressa* and these MTs will be cloned and characterized in the future.

In conclusion, we showed that transcripts encoding three MTs were cloned and sequenced; they encode unique MTs with homology with marine invertebrate and *C. elegans* MTs. UcMTs expressed in bacteria allowed copper and zinc accumulation in vivo. Thus, it is likely that these UcMTs may participate in copper and zinc accumulation in the marine alga *U. compressa*.

### **4. Materials and Methods**

### *4.1. Sampling of Algae and Water Collection*

The green macroalga *U. compressa* was collected during spring 2017 from the high intertidal zone at Cachagüa Beach (32◦34 S), a site in central Chile with no history of metal pollution [23]. Algal samples were transported to the laboratory in sealed plastic bags inside a cool-box. In the laboratory, material was rinsed three times with filtered seawater, cleaned manually, and sonicated three times for 2 min using an ultrasound bath (HiLab Innovation Systems, model SK221OHP) to remove epiphytes. The algae were maintained in aerated seawater under an irradiance of 50 μmoles m−<sup>2</sup> s−<sup>1</sup> on a photoperiod of 12 h:12 h light:dark cycle, at 14 ◦C for 4 days, prior to experimentation. Seawater was obtained from Quintay (33◦12 S), a pristine site in central Chile, filtered through 0.45 and 0.2 μm pore size membranes and stored in darkness at 4 ◦C.

### *4.2. Algal Culture and RNA Extraction*

*U. compressa* (1 g of FT) was cultivated in 100 mL of filtrated seawater containing 10 μM CuCl2 (Merck, Darmstat, Germany) for 3 days. The alga was washed with 10 mM Tris-50 mM EDTA pH 7.0, in order to eliminate copper and other metals bound to cell walls [37].

### *4.3. Purification of Total RNAs and mRNAs for RACE-PCR*

Total RNAs were extracted as described in [38]. *U. compressa* (150 mg of FT) was frozen in liquid nitrogen and pulverized in a mortar. One mL Trizol reagent (Invitrogen, Carlsbad, CA, USA) was added and the alga was homogenized with a pestle until thawing. The mixture was centrifuged at 12,000× *g* for 10 min at 4 ◦C, and the supernatant was recovered. Chloroform (200 μL) was added and the mixture was vortexed for 10 s and left at room temperature for 3 min. The mixture was centrifuged at 12,000× *g* for 15 min at 4 ◦C, and the aqueous phase was recovered. Isopropanol (500 μL) was added and the solution incubated for 10 min at room temperature. The solution was centrifuged at 12,000× *g* for 10 min at 4 ◦C, and the supernatant removed. The pellet was washed twice with 1 mL of 75% ethanol, gently vortexed, and centrifuged at 7000× *g* for 5 min at 4 ◦C. The ethanol phase was removed, the pellet dried for 15 min at room temperature, dissolved in 50 μL of ultrapure water treated with DEPC (water-DEPC), and incubated for 10 min at 60 ◦C.

Total RNAs were quantified using Nanodrop spectrophotometer (Tecan, Zürich, Switzerland); the integrity was verified by agarose gel electrophoresis and stored at −80 ◦C. Messenger RNAs were purified from 100 μg of total RNA using NucleoTrap mRNA minikit (Macherey-Nagel, Düren, Germany), mRNAs were eluted in 25 μL of water-DEPC, and normally 1 μg of mRNAs was obtained from 100 μg of total RNAs.

#### *4.4. Amplification of 5 RACE cDNAs*

The amplification of cDNAs was performed using MMLV reverse transcriptase kit (Promega, Madison, WI, USA). To this end, 4.1 μL of purified mRNAs (150 ng) were mixed with 2.5 μL of primer 1 for MT1 (5 GTAGCAGGCACAGTCGTCA3 ), primer 2 for MT2 (5 AGGCCTAACAAGCAGCGTCC3 ), or primer 3 for MT3 (5 CCGAGAGCGTGTCCTTACTT3 ) at 10 μM, and 8.4 μL of water-DEPC were added to complete a final volume of 15 μL. The mixture was denatured at 65 ◦C for 5 min and cooled on ice for 1 min. Then, 0.7 <sup>μ</sup>L of RNase inhibitor (10 U <sup>μ</sup>L−1), 5 <sup>μ</sup>L of MMLV buffer 5×, 1.5 <sup>μ</sup>L of dNTPs mixture (10 mM of each dNTP), 1 μL de MMLV reverse transcriptase (200 U μL−1), and 1.8 μL of water- DEPC were added to complete a final volume of 25 μL. The mixture was sequentially incubated at 42 ◦C for 50 min, at 50 ◦C for 10 min, at 55 ◦C for 5 min, and at 70 ◦C for 10 min in order to inactivate MMLV reverse transcriptase. cDNAs were treated with RNAse H, purified using GEL/PCR purification mini kit (Favorgen Biotech Corp., Changzhi, Taiwan), and eluted in 40 μL of water-DEPC.

Purified cDNAs (14.5 μL) were incubated with 0.5 μL of terminal transferase (TdT), 20 U μL−1, New England Biolabs, Ipswich, MA, USA), 1 μL of dCTP, 2 μL of CoCl2 (2.5 mM), and 2 μL of TdT buffer 10×, to complete a final volume of 20 μL. The mixture was incubated at 37 ◦C, and at 75 ◦C for 20 min to inactivate TdT in order to obtain cDNAs having a 3 C-tail.

The first round of amplification was performed using PCR kit (Favorgene, London, UK) and 1 μL of C-tailed cDNAs mixed with 10 μL of PCR mixture, 0.6 μL of 5 RACE adapter primer (5 GGCCACGCGTCGACTAGTACGGGIIGGGIIGGGIIG3 ), and with 0.6 μL of primer 4 for MT1 (5 GGCATACGCACGTCTCGGG3 ), primer 5 for MT2 (5 CTGCGTAACGACATAGCCGA3 ), or primer 6 for MT3 (5 GCAGCCAGAATCGCAACTAC3 ), at 10 μM, and 6.8 μL of water-DEPC to complete a final volume of 20 μL. The mixture was incubated at 95 ◦C for 3 min and subjected to 40 cycles of denaturation at 95 ◦C for 5 s, hybridization at 63 ◦C for 10 s and amplification at 72 ◦C for 15 s, using a real-time thermocycler RotorGene 6000. The second round of amplification was performed with the mixture diluted 100 times in distilled water-DEPC, and 14.4 μL of the cDNAs were mixed with 10 μL of PCR mixture, 0.6 μL of abridged universal adaptor primer (AUAP) (5 GGCCACGCGTCGACTAGTAC3 ) and primer 7 for MT1, (5 GCAACCATCTTCGGTTTGGC3 ), primer 8 for MT2, (5 ATCCTTCGCGGGTGAGCAAG3 ), and primer 9 for MT3 (5 CACAGTTGCATTCTGCGGTT3 ) at 10 μM and with 6.8 μL of water-DEPC to complete a final volume of 20 μL. The amplification was performed using 40 cycles of amplification mentioned before. The amplified fragments were analyzed in a 2% agarose gel, stained with SYBR green (Invitrogen, Carlsbad, CA, USA), and visualized on a UV trans-illuminator.

#### *4.5. Amplification of 3 RACE cDNAs*

Initial cDNAs were obtained using MMLV reverse transcriptase kit (Promega, Madison, WI, USA). To this end, 6.8 μL of mRNAs (250 ng) were mixed with 5 μL of 3 RACE adapter primer with an oligo-dT tail (5 GGCCACGCGTCGACTAGTACTTTTTTTTTTTTTTTTT3 ) at 10 μM and with 3.2 μL of water-DEPC to complete a final volume of 15 μL. The mixture was denatured at 65 ◦C for 5 min and then cooled on ice for 1 min. Then, 0.7 <sup>μ</sup>L of RNAse inhibitor (10 U <sup>μ</sup>L−1), 5 <sup>μ</sup>L of MMLV buffer 5×, 1.5 μL of dNTPs mixture (10 mM of each dNTP), 1 μL de MMLV reverse transcriptase (200 U μL−1), and 1.8 μL of water- DEPC were added to complete a final volume of 25 μL. The mixture was incubated

at 42 ◦C for 1 h and at 70 ◦C for 10 min to inactivate MMLV reverse transcriptase and diluted to 50 μL with water-DEPC. cDNAs were treated with RNAse H, purified using GEL/PCR purification mini kit (Favorgen Biotech Corp., Changzhi, Taiwan), and eluted in 40 μL of water-DEPC.

The first round of amplification was performed using PCR kit (Favorgene, London, UK) and 1 μL of cDNAs were mixed with 0.6 μL of AUAP (5 GGCCACGCGTCGACTAGTAC3 ), 0.6 μL of primer 10 for MT1 (5 CAGTGCCAAACCGAAGATGG3 ), primer 11 for MT2 (5 GATGAGGGCTGTCCTTGCTC3 ), or primer 12 for MT3 (5 AGTGTGATGCTGAGTGCTGT3 ) at 10 μM and 6.8 μL of water-DEPC to complete a final volume of 20 μL. The mixture was incubated at 95 ◦C for 3 min and subjected to 40 cycles of denaturation at 95 ◦C for 5 s, hybridization at 63 ◦C for 10 s, and amplification at 72 ◦C for 15 s, using a real-time thermocycler RotorGene 6000. The second round of amplification was performed with the mixture diluted 100 times in distilled water-DEPC, and 14.4 μL of the cDNAs were mixed with 10 μL of PCR mixture, 0.6 μL of AUAP forward (5 GGCCACGCGTCGACTAGTAC3 ) and with primer 13 for MT1 (5 GGTTGCAAGTGCTAGCTGAC3 ), primer 14 for MT2 (5 GCTTGTTAGGCCTCAGTGGT3 ), or primer 15 for MT3 (5 TGTCAGTGCGACAGCCTAA3 ) and with 6.8 μL of water-DEPC to complete a final volume of 20 μL. The amplification was performed using 40 cycles of amplification mentioned before. The amplified fragments were analyzed in a 2% agarose gel, stained with SYBR green (Invitrogen, Carlsbad, CA, USA) and visualized on a UV trans-illuminator.

### *4.6. Amplification of UcMT ORFs*

MT1.1 ORF was amplified using primer 16 forward (5 ATGGACTGCCGTTGCG3 ) and primer 17 reverse (5 GCACTTGCAACCGCCAGAGC3 ); MT2 ORF was amplified using primer 18 forward (5 ATGAACTGCTGTTGCGA3 ) and primer 19 reverse (5 GACACAGCCCGGACAGGC3 ); and MT3 ORF was amplified using primer 20 forward (5 ATGTCGTCTTGTTGTGAAGC3 ) and primer 21 reverse (5 GGCTGTCGCACTGACACAG3 ). The mixture was incubated at 95 ◦C for 2 min and subjected to 35 cycles of denaturation at 95 ◦C for 15 s, hybridization at 65 ◦C for UcMT1.1, 63 ◦C for MT2 and 62 ◦C for UcMT3, during 10 s, and amplification at 72 ◦C for 15 s, using a real-time thermocycler RotorGene 6000.

#### *4.7. Cloning of UcMTs 5 and 3 UTRs, and UcMT ORFs in pGEM-T Vector*

The 5 RACE and 3 RACE amplification fragments obtained in the second PCR (see above) and those of UcMTs ORFs were subjected to electrophoresis in 2% agarose gel. The piece of agarose gel containing the stained fragments was removed from the gel and placed in Eppendorf tubes. The amplified fragments were eluted from agarose using Gel/PCR purification kit (Favorgen, London, UK), recovered in 50 μL of water-DEPC and stored at 4 ◦C. Amplified fragments were ligated with the cloning vector pGEM-T easy (Promega, Madison, WI, USA) and transformed in *E. coli* competent cells One Shot TOP 10 (Invitrogen, Carlsbad, CA, USA). Transformed *E. coli* cells were cultivated in 10 mL of LB medium (10 g tryptone, 5 g yeast extract, and 100 g NaCl in 1 L of distilled water) supplemented with 100 μg mL−<sup>1</sup> of ampicillin. The culture was centrifuged at 3,000 x g for 5 min in a centrifuge model Nuwind (Nuaire, Plymouth, MN, USA). Transformed pGEM-T vectors were purified from the bacterial pellet using Wizard Plus SV Miniprep DNA Purification System (Promega, Madison, WI, USA). To check cloning of 5 UTR fragments in pGEM-T, primers AUAP and primers 7, 8, and 9 were used. To check cloning of 3 UTR in pGEM-T, primer AUAP and primers 13, 14, and 15 were used. PCR conditions were identical to those used for the amplification of 5 and 3 RACE ends (mentioned above). To amplify the ORFs, primers 16 and 17 for MT1, primers 18 and 19 for MT2, and primers 20 and 21 for MT3 were used and to verify the insertion PCR conditions were those used to amplify ORFs (mentioned above). Cloned fragments were sequenced using an ABI3730XL (Macrogen, Seoul, Korea).

### *4.8. Cloning of UcMTs ORFS in pGEX Expression Vector*

*UcMTs* were synthesized and subjected to codon optimization for expression in *E. coli* by Genscript (Piscataway, NJ, USA) and then ligated to the expression vector pGEX-5X-1 (Genscript) which allowed fusion of UcMTs ORFS with the enzyme glutahione-S-transferase (GST) from the platyhelminthes *Schistosoma japonicum* (26 Kda), an enzyme containing a single cysteine that does not bind metals; the fusion proteins were named UcMT1.1-GST, UcMT2-GST, and UcMT3-GST. The recombinant vectors were sequenced by Genscript to verify the correct insertion of complete ORFs.

### *4.9. Transformation of Expression Vectors in Bacteria*

The recombinant expression vectors were transformed in competent *E. coli* strain BL21 (DE3) (Sigma-Aldrich, Saint Louis, MO, USA). To this end, 200 μL of competent cells BL21 (DE3) were incubated with 50 ng of recombinant expression vector containing UcMTs-GST. Then, 800 μL of SOC medium (2% (*w*/*v*) tryptone, 0.5% (*w*/*v*) of yeast extract, 10 mM NaCl, 2.5 mM KCl, 10 mM MgCl2, 10 mm MgSO4, and 20 mM glucose) were added, and cultivated at 37 ◦C for 45 min. An aliquot of 200 μL was cultured on solid LB medium containing 100 μg L−<sup>1</sup> of carbenicillin in a Petri dish overnight. Individual colonies were selected for each UcMT, cultivated with LB medium, and stored at −80 ◦C in LB medium containing 15% glycerol.

### *4.10. Expression of UcMTs-GST in Bacteria*

Recombinant *E. coli* were cultured 15 mL of LB medium until OD600= 0.6, 0.5 mM of isopropyl-β-D-1thiogalactopyranoside (IPTG) was added, and samples of 1 mL were obtained after 1, 3, 6, 9, and 12 h of culture at 37 ◦C. The samples were centrifuged at 7000× *g* for 10 min, washed with PBS pH 7.4 (10 mM Na2HPO4, 1.8 mM KH2PO4, 140 mM NaCl, 2.7 mM KCl), and centrifuged again in similar conditions. Pellets were suspended in 50 μL of protein loading buffer 2× (125 mM Tris HCl pH=6.8, 4% (*w*/*v*) SDS, 20% (*v*/*v*) glycerol, 10% (*v*/*v*) β-mercaptoethanol, 0.004% (*w*/*v*) bromophenol blue), and heated at 95 ◦C for 5 min. A sample of 10 μL was analyzed in a denaturant polyacrylamide gel (12%) and proteins were stained with Coomassie blue staining solution (25% (*v*/*v*) methanol, 5% (*v*/*v*) acetic acid, and 0.1% (*w*/*v*) of Comassie blue G-250).

### *4.11. Purification of UcMTs-GST by A*ffi*nity Column*

A sample of 2 mL of *E. coli* transformed with expression vectors containing a UcMT-GST were added to 100 mL LB medium containing 100 μg mL−<sup>1</sup> ampicillin and cultured at 37 ◦C overnight. A sample of 10 mL was added to 1 L of LB medium containing 100 μg mL−<sup>1</sup> ampicillin and 0.5 mM IPTG, in quadruplicate, until OD600= 0.6 (aprox. 2.5 h). The 4 L of culture were centrifuged at 6000× *g* for 10 min. The pellet was washed twice with 300 mL of PBS and bacteria were suspended in 10 mL of PBS containing 5 mM dithiotreitol (DTT) and 1 tablet of protease inhibitor cocktail (Roche, Manheim, Germany). The bacterial suspension was sonicated for 20 s, with 20 s of pause, for 5 min. The suspension was centrifuged at 6000 rpm for 10 min and supernatant was recovered. Protein concentration was determined using Bradford method [39] and adjusted with PBS-5 mM DTT (PBS-DTT) to 1 mg mL<sup>−</sup>1. UcMTs-GST were purified by HPLC using GSTrap HP (General Electric, Uppsala, Sweden) at 5 bars of pressure, washed with PBS-DTT, and eluted with 3 mL of buffer 50 mL Tris-HCl-10 mM GSH. Normally, 1–2 mg of purified UcMT-GST was obtained from 4 L of bacterial culture and proteins were quantified using Bradford method [40].

### *4.12. Detection of UcMTs-GST with Anti-GST Antibody*

Transformed bacteria were cultured in 100 mL of LB medium until OD600 = 0.6, 0.5 mM IPTG was added and the mixture incubated for 6 h. The culture was centrifuged at 6000× *g* for 5 min, the pellet suspended in 5 mL of buffer PBS, and sonicated for 20 s, with 20 s of pause, for 5 min. Proteins (20 μg) were separated by electrophoresis in a denaturant 12% polyacrylamide gel and transferred to a nitrocellulose membrane for 10 min using Trans Blot Turbo apparatus (BioRad, Hercules, CA, USA). The membrane was stained with Ponceau Red dye and washed with 10 mL of distilled water. The membrane was incubated in 10 mL TTBS (20 mM Tris-HCl pH 7.5, 0.1 mM NaCl, 0.05% Tween-20) containing 5% skim milk for 1 h, washed twice with 10 mL TTBS for 15 min, incubated with 10 mL TTBS

containing 3% skim milk and the antibody anti-GST (Sigma-Aldrich, St Louis, MO, USA) diluted 5000 times, and washed four times in TTBS for 15 min. The membrane was incubated in TTBS containing 3% skim milk and the secondary antibody prepared against rabbit IgG coupled to hoseradish peroxidase (Agrisera, Vännas, Sweden) diluted 2000 times, for 1 h, and washed four times with TTBS for 15 min. Proteins were detected using ECL Western Blotting System kit (Amersham, Buckinghamshire, UK) and revealed using a C-Digits chemiluminiscence Western blot scanner Li-Cor (Lincoln, NE, USA) and Image Studio Digits software version 4.0 Li-Cor.

### *4.13. Quantification of Copper and Zinc in Bacteria Expressing UcMTs-GST*

Recombinant bacteria were cultured in 100 mL of LB medium containing 100 mg mL−<sup>1</sup> of carbenecillin until DO600 = 0.6, with 0.5 mM IPTG for 30 min, and with 1 mM CuSO4 or 1 mM ZnCl2 and IPTG for 6 h. Bacterial pellets showed a weight of 26–42 mg for copper cultures and 46–62 mg for zinc cultures. Pellets were dried at 60 ◦C for 48 h, suspended in 5 mL of 60% (*v*/*v*) HNO3, and incubated at 85 ◦C for 2 h. The solutions were filtered through 0.22 μm MCE filters (TCL, Santiago, Chile) and analyzed by flame atomic emission spectrophotometry using an atomic emission spectrophotometer ThermoFisher (Waltham, MA, USA).

### *4.14. Hierarchical Clustering of UcMTs*

Amino acid sequences corresponding to MTs of different animal and plant species (234 in total) were selected from revised SwissProt repository of the UniprotKB database (www.uniprot.org). Alignment of these sequences was performed with Clustal W software with default setting. This alignment was used to generate the phylogenetic reconstruction to represent a hierarchical clustering using UPGMA algorithm based on distance. Phylogenetic and hierarchical clustering analyses were conducted using MEGA software version X [39].

### *4.15. Statistical Analyses*

Statistical analyses were performed with the Prism 6 statistical package (Graph Pad software Inc., San Diego, CA, USA). Following confirmation of normality and homogeneity of variance, significant differences among treatments were determined by two-way ANOVA and Tukey's multiple comparison post-hoc test, at a 95% confidence interval.

### **Supplementary Materials:** Supplementary materials can be found at http://www.mdpi.com/1422-0067/21/1/153/s1.

**Author Contributions:** A.Z. amplified and cloned 5 and 3 UTR and ORFs of transcripts encoding UcMTs, expressed UcMTs-GST in bacteria, and analyzed copper and zinc accumulation. A.G. and M.G. purified UcMTs-GST. D.L. guided A.Z., did hierarchical clustering of MTs, Western hybridization of UcMTs-GST, and UV-VIS spectra of MTs-GST. C.A.S. revised the manuscript and participate in discussions, and A.M. designed experimental work and wrote the manuscript. All authors have read and agreed to the published version of the manuscript.

**Funding:** This work was financed by Fondecyt 1160013 to A.M.

**Acknowledgments:** We thank Luis Lemus and Cristián Vera who helped with UV-VIS spectra analyses.

**Conflicts of Interest:** The authors declare no conflict of interest.

### **Abbreviations**


### **References**


© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Review* **Advances in the Mechanisms of Plant Tolerance to Manganese Toxicity**

**Jifu Li 1,2, Yidan Jia 1,2, Rongshu Dong 1,3, Rui Huang 1,3, Pandao Liu 1,3, Xinyong Li 1,3, Zhiyong Wang 2, Guodao Liu 1,3 and Zhijian Chen 1,2,3,\***


Received: 22 September 2019; Accepted: 12 October 2019; Published: 14 October 2019

**Abstract:** Manganese (Mn) is an essential element for plant growth due to its participation in a series of physiological and metabolic processes. Mn is also considered a heavy metal that causes phytotoxicity when present in excess, disrupting photosynthesis and enzyme activity in plants. Thus, Mn toxicity is a major constraint limiting plant growth and production, especially in acid soils. To cope with Mn toxicity, plants have evolved a wide range of adaptive strategies to improve their growth under this stress. Mn tolerance mechanisms include activation of the antioxidant system, regulation of Mn uptake and homeostasis, and compartmentalization of Mn into subcellular compartments (e.g., vacuoles, endoplasmic reticulum, Golgi apparatus, and cell walls). In this regard, numerous genes are involved in specific pathways controlling Mn detoxification. Here, we summarize the recent advances in the mechanisms of Mn toxicity tolerance in plants and highlight the roles of genes responsible for Mn uptake, translocation, and distribution, contributing to Mn detoxification. We hope this review will provide a comprehensive understanding of the adaptive strategies of plants to Mn toxicity through gene regulation, which will aid in breeding crop varieties with Mn tolerance via genetic improvement approaches, enhancing the yield and quality of crops.

**Keywords:** manganese toxicity; Mn detoxification; tolerance mechanism; gene function; subcellular compartment

### **1. Introduction**

Manganese (Mn) is the second most prevalent trace element in the Earth's crust after iron (Fe), and is widely distributed in soils, sediments, and other biological materials [1]. In soils, Mn is present in a wide range of oxidation states, including Mn(II), Mn(III), Mn(IV), Mn(VI), and Mn(VII) [2]. Among the oxidized forms of Mn, divalent Mn(II) is the most soluble species in soils and is also the most available form of Mn for plant acquisition. The solubility of Mn is strongly influenced by soil pH and redox conditions [1,3]. At neutral or higher soil pH, Mn(III) and Mn(IV) are the predominant and insoluble forms of Mn. However, in poorly drained acid soils with pH levels below 5.0 and a reducing environment, oxidized Mn is easily reduced to divalent Mn [4]. Thus, the available Mn in soils is variable and generally ranges from 450 to 4000 mg per kilogram [3]. For example, the concentration of Mn varies between 40 and 1681 mg per kilogram in farmland soils across mainland China [5], while the concentration of Mn in the agricultural soils of central Greece is from 685 to 1307 mg per kilogram [6].

Mn is an example of a transition element that is required for humans, animals, and plants. For most plants, Mn is absolutely necessary at low levels of 20–40 mg per kilogram dry weight [7,8]. Mn is involved in a variety of metabolic processes, including photosynthesis, respiration, fatty acid and protein synthesis, as well as enzyme activation. For example, Mn is an indispensable constitutive element in the Mn cluster structure of the oxygen-evolving complex in photosystem II (PSII) that participates in the water-splitting process, providing necessary electrons for photosynthesis [9,10]. Mn acts as an important cofactor of various enzymes, including superoxide dismutase (MnSOD), catalase (MnCAT), decarboxylases of the tricarboxylic acid (TCA) cycle, and RNA polymerases [8,11]. In addition, Mn is required for multiple steps in the biosynthesis of secondary metabolites, such as lignins, flavonoids, cinnamic acid, and acyl lipids [12].

Despite its necessity, Mn is also considered one of the heavy metals that can be harmful to plants at excessive levels. When the Mn concentration in the aboveground tissues of plants reaches 150 mg per kilogram dry weight, Mn toxicity can generally occur, especially for plants growing in acid soils [13,14]. Many previous studies demonstrate that Mn toxicity can disrupt various physiological processes in plant cells, such as triggering oxidative stress, inhibiting enzyme activity, impeding chlorophyll biosynthesis and photosynthesis, and preventing the uptake and translocation of other mineral elements, including phosphorus (P), Fe, and magnesium (Mg) [14–16]. As a result, Mn toxicity leads to the appearance of toxicity symptoms, including chlorosis in young leaves, necrotic dark spots on mature leaves, and crinkled leaves, ultimately inhibiting plant growth. Symptoms of Mn toxicity vary widely among plant species and varieties. For example, chlorosis and necrosis have been reported in leaves of common bean (*Phaseolus vulgaris*) [17], clover (*Trifolium repens*) [18], ryegrass (*Lolium perenne*) [19], and stylo (*Stylosanthes guianensis*) [20]. Brown spots surrounded by irregular areas of chlorotic tissues are observed in cowpea (*Vigna unguiculata*) [21], soybean (*Glycine max*) [22], and barley (*Hordeum vulgare*) [23]. The diverse expressions of Mn toxicity probably indicate different Mn-tolerant capabilities among plant species and cultivars. For example, among different legumes, *Medicago sativa*, *Trifolium fragifevum*, *Leucaena leucocephala*, and *Medicago tvuncatula* are considered the most sensitive to Mn toxicity, while *Centrosemapubescens*, *Lotononis bainesii*, Townsville stylo (*Stylosanthes humilis*), and *Desmodium mcinatum* are the most tolerant plant species [24].

Over the last few decades, there have been major advances in elucidating the mechanisms underlying plant tolerance to Mn toxicity at multiple levels, from physiological changes to biochemical responses (Figure 1). For example, activation of the antioxidant system, including the free radical-mitigating antioxidant enzymes and nonenzymatic components, is thought to be vital for plants alleviating excess Mn-induced oxidative stress [25]. The important roles of the regulation of Mn uptake, translocation, and distribution have been implicated in many plants' responses to Mn toxicity, such as rice (*Oryza sativa*) [26,27], Arabidopsis (*Arabidopsis thaliana*) [28], and Caribbean stylo (*Stylosanthes hamata*) [29]. Furthermore, plants can sequester Mn into subcellular compartments, such as vacuoles, the endoplasmic reticulum (ER), Golgi apparatuses, and cell walls, to withstand the toxic effects of high Mn stress [30,31]. In addition, free Mn ions can be chelated with protein-based, organic, and inorganic compounds to form inactive Mn complexes, combating the deleterious effects of Mn toxicity [18–20].

To date, a variety of genes and proteins have been shown to be involved in the responses to Mn toxicity of plants, such as orange (*Citrus sinensis*) [32], common bean [33], tomato (*Solanum lycopersicum*) [34], stylo [20,35], cowpea [21,36], soybean [22], rice, and barley [23]. Many of the identified genes have been functionally integrated into specific pathways, illuminating the molecular processes of the plant response to Mn toxicity. Furthermore, the functions of numerous genes involved in Mn detoxification through regulation of Mn uptake, distribution, and accumulation have been well characterized in plants [29,37–39]. Therefore, the purpose of this review is mainly to focus on Mn as a toxic transition metal to plants and the mechanisms of plant tolerance to Mn stress. This review will discuss the current understanding of plant genes involved in Mn uptake, distribution, and accumulation, which contribute to Mn detoxification. Furthermore, we also highlight

the candidate genes that can potentially be used for breeding crop varieties tolerant to Mn toxicity via genetic improvement approaches.

**Figure 1.** Schematic representation of Mn toxicity and strategies for increasing Mn tolerance in plants. Mn toxicity can trigger oxidative stress and disrupt photosynthesis, which may result in the generation of interveinal chlorosis in young leaves, necrotic dark spots on mature leaves, and crinkled leaf. Furthermore, Mn toxicity can lead to the formation of brown roots and prevent the uptake and translocation of other mineral elements. In plants, Mn tolerance strategies include modification of Mn translocation and distribution, sequestration of Mn into subcellular compartments, modulation of the antioxidant system, changes in biochemical pathways, and regulation of Mn transporters. In addition, the mediation of root exudates, the application of Si in roots, and the amelioration of soil acidification, humidity, and organic matter content also contribute to increase plant Mn tolerance. Red arrows indicate the toxic effects of excess Mn to plants. Purple arrows represent the adaptive strategies of plants to Mn toxicity.

### **2. Activation of the Antioxidant System**

As a toxic metal, excess Mn can generate reactive oxygen species (ROS) and trigger oxidative stress in plants, causing lipid peroxidation and damaging photosynthetic pigments and proteins if ROS are not well scavenged [25,35]. One of the adaptive changes that alleviates the toxic effects of high Mn in plants involves the activation of the antioxidant system via antioxidant enzymes, such as superoxide dismutase (SOD), peroxidase (POD), catalase (CAT), ascorbate peroxidase (APX), and glutathione reductase (GR), and nonenzymatic antioxidant components, including ascorbate (AsA) and glutathione (GSH) [35,40]. Increases in the activities of antioxidant enzymes under Mn toxicity are generally associated with enhanced Mn tolerance in common bean [41], cucumber (*Cucumis sativus*) [42], and perennial ryegrass [40]. In perennial ryegrass, for example, the Mn-tolerant ryegrass cultivar Kingston exhibits higher SOD activity than the Mn-sensitive ryegrass cultivar Nui—a higher expression of the *Fe–SOD* gene is observed in Kingston compared to that in Nui [40]. Thus, the induced *Fe–SOD* expression in Kingston is likely to contribute to its high Mn-toxicity tolerance. Additional studies in cowpea have shown that both the activities of H2O2-producing and H2O2-consuming PODs are enhanced by Mn toxicity in the leaf apoplast [21]. Furthermore, proteomic analysis indicated that the protein accumulation of PODs in the leaf apoplast is increased by high Mn [21]. Similar results have been implicated in citrus and stylo, in which the expression of *POD* genes is enhanced when plants are subjected to Mn toxicity [35,43]. Therefore, it is probable that SOD and POD represent two key proteins in the plant defense against oxidative damage caused by Mn toxicity. However, considering the damage caused by Mn toxicity, ROS-scavenging systems, through regulation of the antioxidant system, seem to be insufficient to alleviate oxidative stress, which might be a general response of plants to Mn toxicity.

### **3. Regulation of Mn Uptake**

Although Mn is required in relatively small amounts, the Mn content accumulated in most plants is approximately 30–500 mg per kilogram dry weight, which is higher than their normal growth requirements [8,14,44]. Therefore, it is reasonable to propose that there are some key transporter genes responsible for Mn acquisition in response to high Mn stress (Figure 2). Studying the mechanisms of plant Mn transport can greatly increase our understanding of how plants acquire and transport Mn under variable environmental Mn levels.

**Figure 2.** Summary of genes affecting Mn transport and tolerance in plants. Squares: Import into the cytosol; circles: Export out of the cytosol; blue: MTP family; green: ZTP family; red: Nramp family; purple: YSL family; yellow: IRT family; orange: ECA family; cyan: CAX family; brown: DMT family; gray: unknown. ER: Endoplasmic reticulum; Nramp: Natural resistance-associated macrophage protein; MTP: Metal tolerance protein; DMT: Divalent metal transporter; ZIP/IRT: Zinc-regulated transporter/iron-regulated transporter-like proteins; YSL: Yellow stripe-like protein; CAX: Cation exchanger; ECAs: ER-type calcium ATPases; MDH: Malate dehydrogenase; ALMT: Aluminum-activated malate transporter; OAA: Oxaloacetate; ROS: Reactive oxygen species; SOD: Superoxide dismutase; POD: Peroxidase; CAT: Catalase. At: *Arabidopsis thaliana*; Os: *Oryza sativa*; Gm: *Glycine max*; Hv: *Hordeum vulgare*; Mt: *Medicago tvuncatula*; Cs: *Cucumis sativus*; Sh: *Stylosanthes hamata*; Sg: *Stylosanthes guianensis*; Le: *Lycopersicon esculentum*; Bm: *Beta vulgaris* subspecies maritima; Pt: *Populus trichocarpa*. Question marks behind some genes mean that the exact roles of these genes or their localizations remain to be further clarified.

The major transporters responsible for Mn acquisition in plants are members of the natural resistance-associated macrophage protein (Nramp) family, which have so far been functionally characterized in many plants, for example, AtNramp1 from Arabidopsis, OsNramp5 from rice, and HvNramp5 from barley [26,45]. In Arabidopsis, AtNramp1, belonging to the Nramp family, is the major high-affinity Mn transporter involved in Mn uptake. AtNramp1 is localized to the plasma membrane. The transcripts of *AtNramp1* are mainly detected in roots, where their levels are ten times greater than in shoots. Furthermore, *AtNramp1* transcripts are increased by Mn deficiency in the roots [46]. AtNramp1 can complement the phenotype of a yeast mutant, *smf1*, which is defective in Mn uptake when grown in medium containing the divalent cation chelator EGTA [47]. Furthermore, when cultivated in a medium lacking Mn, the T-DNA insertion mutant Atnramp1-1 produces less biomass than wild-type Arabidopsis. The growth inhibition of the mutant can be attributed to less Mn accumulation compared to the wild-type plants under Mn-deficient conditions [46].

In rice, Mn uptake is mediated by OsNramp5, a homolog of AtNramp1 [48]. In contrast to Arabidopsis, OsNramp5 is constitutively expressed in roots, and its expression is enhanced by Fe and zinc (Zn) deficiency but does not respond to different Mn levels in roots [48]. As OsNramp5 can complement the growth of yeast strains defective in Mn and Fe transport, OsNramp5 is implicated in Mn and Fe transport [49]. As OsNramp5 is polarly located at the distal side of both the exodermis and endodermis of mature roots, OsNramp5 is likely to act as an influx transporter and acquire Mn from the soil to the exodermal cells as well as from the apoplastic solution to endodermal cells [48]. Knockout of *OsNramp5* resulted in a decreased concentration of Mn and Fe but not Zn in the shoots, suggesting that OsNramp5 is able to transport Fe in addition to Mn. However, the growth of *OsNramp5* knockout lines is unaffected when the Fe concentration in the external solution is decreased, and the Fe concentrations in the shoots and roots are similar to those of the wild type under Fe deficiency. Thus, the authors conclude that the uptake of Fe required for growth is mediated by other transporters, and OsNramp5 is responsible for additional Fe uptake [48]. A similar key role has been assigned to metal tolerance protein 9 (OsMTP9), the other type of transporter belonging to the cation diffusion facilitator (CDF) family that participates in Mn uptake and translocation in rice roots [50]. *OsMTP9* shows higher expression in roots, but its expression is not influenced by external Mn levels [50]. Tissueand cell-specific localization analysis revealed that OsMTP9 is localized to the proximal sides of both the exodermis and endodermis of mature root zones, which is opposite to the sites of of OsNramp5 localization in rice roots. Further evidence shows that OsMTP9 acts as an efflux transporter and is responsible for Mn translocation to the root stele [50]. Therefore, the different polar localizations of OsNramp5 and OsMTP9 facilitate Mn uptake from the soil solution to the stele in rice.

Similar results have also been found for HvNramp5, which is localized to the plasma membranes of the epidermal cells of the root tips in the outer root cell layers of barley [51]. There is evidence that HvNramp5 displays transport activity for both Mn and cadmium (Cd) when expressed in yeast cells, and disruption of *HvNramp5* results in growth reduction in barley under low Mn supply [51]. Therefore, HvNramp5 is a transporter required for Mn uptake in barley. In addition, GmDMT1 (divalent metal transporter 1), a nodule-enhanced transporter belonging to the Nramp family in soybean, has also been found to transport Mn in addition to Fe when expressed in yeast [52], although further investigation is needed to understand the physiological roles of GmDMT1 in Mn acquisition in soybean. In addition, members of the zinc-regulated transporter/iron-regulated transporter-like proteins (ZRT/IRT) family were found to have the ability to transport Mn, such as HvIRT1 from barley [53].

Considering the particular importance of the transporter genes controlling Mn uptake in plants, it is reasonable to propose that increased Mn detoxification can be achieved through decreased Mn accumulation from decreasing excess Mn uptake and root-to-shoot Mn translocation, by downregulating transporter genes specific for Mn uptake under high Mn stress. Therefore, manipulation of these transporter genes is an alternative strategy to facilitate the plant response to varying Mn levels through regulation of Mn acquisition.

### **4. Regulation of Mn Translocation and Distribution**

After Mn is taken up by roots, most Mn is translocated from roots to shoots and further delivered to various tissues for growth requirements. Thus, it is important to understand the long-distance and whole-plant translocation of Mn in plants in response to different Mn levels, from limited to excessive. In Arabidopsis, two ZIP members, AtZIP1 and AtZIP2, are implicated in Mn translocation from roots to shoots [54]. Both AtZIP1 and AtZIP2 are mainly expressed in the root stele and do not respond to external Mn levels at the transcriptional level. AtZIP1 and AtZIP2 localize to the tonoplast and plasma membrane, respectively. It is probable that AtZIP1 functions in the remobilization of Mn from vacuoles to the cytoplasm in root stellar cells, while AtZIP2 plays a role in Mn movement to the root vasculature for further translocation to the shoots [54]. The loss-of-function mutants of the *AtZIP1* gene in Arabidopsis show severe sensitivity to Mn deficiency. However, the T-DNA *AtZIP2* knockout lines display more tolerance to Mn toxicity than the wild type [54]. Furthermore, Mn concentration in the roots of *AtZIP2* knockout lines is much higher than that in wild-type plants, but no significant differences in shoot Mn concentrations are observed between knockout lines and wild-type plants [54]. Considering that AtZIP2 has high root expression in the stele, AtZIP2 is likely to play a role in Mn transport into the root vasculature, which ultimately helps to provide Mn to the xylem parenchyma, where other transporters such as the heavy metal ATPase, AtHMA2/4, may mediate xylem loading of Mn to the shoot in the transpiration stream as proposed by the authors [54].

OsYSL2, belonging to the yellow stripe-like family, has been characterized to function in long-distance Mn transport and distribution in rice [55]. *OsYSL2* is mainly expressed in leaves, flowers, and developing seeds [55,56]. Electrophysiological measurements using *Xenopus laevis* oocytes show that OsYSL2 is involved in the transportation of Mn–nicotianamine (NA) in addition to Fe–NA complexes [56]. The phloem and seed localization of OsYSL2 suggests that OsYSL2 transports Mn–NA and Fe–NA complexes via the phloem and then loads these complexes into the grain [56]. Overexpression of *OsYSL2* leads to increases in Mn accumulation in the grain [55], suggesting that OsYSL2 is involved in the translocation of Mn into the grain. In addition, some evidence suggests that Mn complexes may be delivered by other transporters, such as *AtOPT3* (a putative oligopeptide transporter) and *AtYSLs* from Arabidopsis [57–59], and *ZmYS1* from maize [60], but the exact roles of these genes remain to be clarified.

Additional studies have shown that rice OsNramp3 is a plasma membrane-localized influx transporter for the distribution of Mn, but not Fe and Cd [37]. *OsNramp3* displays higher expression in the nodes and is not affected by external Mn at the transcriptional level. It is noteworthy that the OsNramp3 protein is rapidly degraded within a few hours when plants are exposed to high Mn stress [37]. OsNramp3 is proposed to function with the following patterns: Under Mn deficiency, OsNramp3 preferentially transports Mn to young leaves and panicles via intervascular transfer, but in contrast, under excess Mn conditions, due to rapid OsNramp3 protein degradation, Mn is delivered to old tissues, protecting developing tissues from the toxic effects of excess Mn [37]. Therefore, the authors suggest that OsNramp3 functions as a node-based switch for Mn distribution, which turns the protein on or off in response to variable environmental Mn levels. These findings above provide a major advancement in the understanding of Mn distribution in plants through the regulation of transporters at the post-translational level.

### **5. Intracellular Mn Detoxification in Subcellular Compartments**

As the amount of Mn accumulated in most plants usually exceeds their normal growth requirements, plants must cope with excess Mn via internal detoxification. In this regard, one of the key strategies for plant tolerance to Mn toxicity is the compartmentalization of Mn into subcellular compartments [14]. Therefore, transporters that localize to the endomembrane compartments are suggested to be critical for intracellular Mn detoxification in plant cells.

The vacuole, an organelle that comprises approximately 90% of the total cell volume, is the dominant sink for various toxic compounds, including Mn [61]. Some transporters belonging to the CDF family act as proton antiporters for efflux metals (e.g., Zn, Fe, Mn, and Cd) out of the cytoplasm or into subcellular compartments (e.g., vacuoles) [62]. ShMTP1, the first functionally characterized CDF for Mn transport into the vacuoles, was isolated from Caribbean stylo, a tropical legume with superior Mn tolerance [29,35,45]. Evidence shows that ShMTP1 is localized to the tonoplast, and overexpression of *ShMTP1* confers Mn tolerance in yeast cells and Arabidopsis via sequestration of Mn into the vacuoles [29]. In addition to ShMTP1, other CDF members, such as OsMTP8.1, also participate in delivering Mn to vacuoles for Mn sequestration [63]. The transcript of *OsMTP8.1* is mainly detected in shoots and is enhanced by high Mn levels. OsMTP8.1 is expressed in all cells of leaf blades and is also localized to the tonoplast. In rice, knockout of *OsMTP8.1* results in the generation of symptoms of Mn toxicity when plants are exposed to high Mn toxicity [63]. However, OsMTP8.1 is not a unique CDF in mediating Mn transport into vacuoles in rice. OsMTP8.2, a homolog of OsMTP8.1, is also involved in Mn sequestration, and loss of function of *OsMTP8.2* results in severe growth inhibition of both shoots and roots of the *osmtp8.1* mutant in the presence of high Mn [64]. Therefore, it is probable that OsMTP8.2 mediates Mn tolerance together with OsMTP8.1 by sequestering Mn into vacuoles. To date, a set of *MTP* homolog genes have been characterized with similar functions in sequestering Mn into vacuoles, such as *AtMTP8* from Arabidopsis [37], *CsMTP8*/*9* from cucumber [65,66], and *CsMTP8* from the tea plant (*Camellia sinensis*) [67]. The conserved function of MTPs among different plant species fully supports the dominant roles of MTPs in Mn detoxification.

Another major transporter for intracellular Mn sequestration into vacuoles is a member of the cation exchanger (CAX) family with metal/H<sup>+</sup> antiport activity. In Arabidopsis, the role of AtCAX2 in Mn transport was confirmed by its ability to confer tolerance to Mn toxicity when its expression was heterologous in *pmc1vcx1cnb*, a Mn-sensitive yeast mutant. A three-amino acid Mn-binding region (Cys–Ala–Phe) in AtCAX2 was subsequently found to be critical for Mn-transport activity [68–70]. Further analysis showed that overexpression of *AtCAX2* in tobacco (*Nicotiana tabacum*) increases the resistance to Mn toxicity via mediating the sequestration of Mn into the vacuoles [68]. In addition to AtCAX2, AtCAX4 and AtCAX5, which localize to the vacuolar membrane, also display Mn2+/H<sup>+</sup> antiport activity [71,72]. The transcripts of both *AtCAX4* and *AtCAX5* in roots are increased under conditions of high Mn [71–73]. Phenotypic analysis shows that transgenic tobacco overexpressing *AtCAX4* displays tolerance to Mn toxicity, while *AtCAX5* can rescue the growth of Mn-sensitive yeast, suggesting their roles in conferring Mn tolerance [72,74]. Arabidopsis mutants, including *cax1*, *cax2*, *cax3*, *cax1*/*cax2,* and *cax2*/*cax3*, have been generated and analyzed for their growth performances under excess Mn levels. Among these mutants, *cax2* and *cax2*/*cax3* displayed severe sensitivity to high Mn stress [75].

An alternate mechanism of intracellular-Mn tolerance in plants is the sequestration of Mn into the Golgi apparatus or endoplasmic reticulum (ER) [15]. AtMTP11 is suggested to be involved in this process in Arabidopsis. AtMTP11 can rescue the growth of yeast mutant *pmr1*, which is defective in a Ca<sup>2</sup>+/Mn2+–ATPase, in the presence of excess Mn. Arabidopsis mutants impaired in *AtMTP11* are sensitive to high Mn levels, whereas plants overexpressing *AtMTP11* are more tolerant to Mn toxicity [76]. In contrast to ShMTP1, OsMTP8.1, and OsMTP8.2 mentioned above, AtMTP11 is localized to a punctate endomembrane compartment probably in the trans-Golgi, but not to the plasma membrane and vacuole. Therefore, a secretory pathway involving vesicular trafficking and exocytosis mediated by AtMTP11 is believed to help increase Mn tolerance in Arabidopsis [28]. Similar functions of other MTPs in sequestering Mn into the Golgi apparatus have been reported for OsMTP11 from rice [27], HvMTP8.1 and HvMTP8.2 from barley [77], PtMTP11.1 and PtMTP11.2 from poplar (*Populus trichocarpa*) [28], as well as BmMTP10 and BmMTP11 from beets (*Beta vulgaris*) [78].

It has been well demonstrated that ER-type calcium ATPases (ECAs), belonging to the Ca2<sup>+</sup>–ATPase subfamily, can use energy from ATP hydrolysis to catalyze the translocation of cations across membranes [79,80]. There are four predicted ECAs in Arabidopsis (AtECA1–4) and three in rice (OsECA1–3) [79]. In Arabidopsis, AtECA1 and AtECA3 are localized to the ER and Golgi compartments, respectively [81–83]. The expression of AtECA1 and AtECA3 was found in all major organs of

Arabidopsis, especially in the roots [81,83]. Both AtECA1 and AtECA3 are able to rescue the growth of yeast under high Mn stress [81,84]. Furthermore, under excess-Mn conditions, the Arabidopsis *ateca1-1* mutants display inhibited root growth, and the growth of the *ateca1-1* mutant is rescued by overexpression of *AtECA1* [81]. Similarly, the root growth of the *ateca3* mutant is impaired by excess Mn, confirming that AtECA3 is also necessary for Mn detoxification in Arabidopsis [84]. Therefore, AtECA1 and AtECA3 are the two key components required for delivering Mn into the ER and Golgi compartments for Mn tolerance. In addition, the YSL family is also implicated in the sequestration of Mn into endomembrane compartments. AtYSL4 and AtYSL6 are reported to be localized to vacuole membranes and internal membranes resembling the ER in Arabidopsis. Significant decreases in fresh weight have been observed in single mutants and double mutants of *AtYSL4* and *AtYSL6* compared to wild-type Arabidopsis grown in high Mn for 21 d [59]. The authors suggest a role for AtYSL4 and AtYSL6 in the sequestration or efflux of this metal into intracellular compartments [59]. However, future characterization of YSL as well as ECAs in other crop species is needed to confirm their exact roles in Mn detoxification via sequestration of Mn into intracellular compartments.

OsYSL6 is reported to transport Mn from the apoplast to the symplast, which is required for the detoxification of excess Mn in rice [84]. Although the expression of *OsYSL6* does not respond to either deficiency or toxicity of Mn, ectopic expression of *OsYSL6* in the yeast mutant indicates transport activity for the Mn–NA complex. Furthermore, knockout of *OsYSL6* in rice increases Mn accumulation in the leaf apoplast but not in the symplast under high Mn stress, resulting in the development of necrosis in the old leaves, a symptom of Mn toxicity [84]. As divalent Mn accumulated in the apoplast can potentially be oxidized to trivalent Mn, which further oxidizes proteins and lipids, causing deleterious effects of Mn toxicity [21], OsYSL6 is likely to alleviate excess Mn toxicity via the transport of Mn from the apoplast to the symplast in rice.

Most of the Mn transporter genes mentioned above display no or only slight responses to varying Mn levels, which may partially explain why plants accumulate large amounts of Mn that far exceed their growth requirements. Therefore, it is of great importance to investigate the regulatory mechanisms of the plant response to external Mn in the future.

### **6. Si Application Alleviates Mn Toxicity**

Another strategy for increasing Mn tolerance can be achieved by the application of silicon (Si) to the roots of plants such as rice [85], cowpea [86,87], and cucumber [88]. The mechanisms for Si-alleviated Mn toxicity include decreasing the Mn accumulation in shoots, promoting Mn oxidation in roots and increasing the cell wall-binding capacity for Mn [88–90]. A recent study showed that supplementation with Si successfully decreases the Mn concentration in the shoots but increases Mn in the roots of rice under high Mn stress, alleviating Mn toxicity [90]. However, Si application cannot alleviate Mn toxicity in the rice *lsi1* mutant, which is defective in Si uptake. OsLsi1 is a Si transporter that transports Si from the external solution to the root cells in rice [91]. Interestingly, the expression of *OsNramp5* is decreased by long-term exposure to Si in the wild type but not in the *lsi1* mutant. The authors suggest that the Si-alleviated Mn toxicity in rice can be attributed to inhibition of root-to-shoot translocation of Mn and decreased Mn uptake by downregulation of Mn transporters, such as OsNramp5 and OsMTP9 [90]. Therefore, OsLsi1 might participate in Mn detoxification through regulation of Si uptake, which deserves further clarification.

### **7. Organic Acid Mediates Mn Detoxification**

Mn can be chelated with protein-based, organic, and inorganic compounds to form Mn complexes, thus decreasing Mn uptake and/or Mn phytotoxicity. Regulation of organic acid metabolism is an important strategy in Mn detoxification. Intracellular Mn in cowpea, *Gossia bidwillii*, and *Phytolacca acinosa* is found to be chelated in complexes with internal citrate, malate, and oxalate, respectively [92–94]. The complexation of Mn by organic acids in the apoplast is proposed to decrease Mn phytotoxicity in cowpea [87]. Increases in internal malate concentrations are observed in leaves and roots of

the Mn-tolerant stylo genotype Fine-stem under high Mn stress, and are closely linked to its Mn tolerance capabilities [20]. Accordingly, Mn might be chelated by malate to form Mn–malate complexes, ultimately conferring Mn tolerance in stylo. Subsequent analysis shows that malate synthesis in stylo could be attributed to a Mn-enhanced malate dehydrogenase (SgMDH1), which catalyzes the reversible conversion of oxaloacetate to malate. Due to successful increases in resistance to Mn toxicity in both yeast cells and Arabidopsis, SgMDH1 is hypothesized to be involved in Mn detoxification through mediated malate synthesis [20].

On the other hand, increases in organic acid exudation from roots in response to Mn toxicity are found in stylo, clover, and ryegrass [18–20]. Increased root exudates of oxalate and citrate in Mn-tolerant ryegrass cultivars have been implicated in increasing Mn tolerance by decreasing Mn uptake from the rhizosphere [19]. Similar results are also reported in stylo, where increased malate exudation from roots helps to confer Mn tolerance, and exogenous malate application to the growth medium increases the resistance of the Mn-sensitive stylo genotype to the toxic effects of Mn [20]. Interestingly, the expression of an aluminum-activated malate transporter (*SgALMT1*) is enhanced by high Mn stress in the Mn-tolerant stylo genotype [20], which likely functions in mediating malate efflux from roots, as observed in aluminum detoxification [95]. Therefore, it is reasonable to hypothesize that coordinated regulation of malate synthesis and exudation by *SgMDH1* combined with *SgALMT1* might facilitate the tolerance of stylo to Mn toxicity.

### **8. Other Aspects**

In recent years, the development of biotechnologies, such as RNA-seq and proteomics, has provided favorable platforms to reveal complex responses of plants to biotic and abiotic stresses [35,96,97]. Many differentially expressed genes and proteins have been previously identified in plants' responses to Mn toxicity. For example, various Mn-responsive genes have been isolated from leaves of citrus using cDNA–AFLP technology, and the identified genes can be classified into different functional categories, such as biological regulation and signal transduction (e.g., protein phosphatase 2a and Myb family transcription factor), carbohydrate and energy metabolism (e.g., ATP synthase subunit alpha and UDP-glycosyltransferases), nucleic acid metabolism (e.g., DNA polymerase phi subunit and histone H4), protein metabolism (e.g., ribosomal proteins, eukaryotic initiation factors, and glutathione S-transferase Tau2), cell wall metabolism (e.g., cell wall-associated hydrolase and glycoside hydrolase family 28 protein), stress responses (e.g., *CAT*, *POD42*, and monodehydroascorbate reductase), and cell transport (e.g., ABC transporter family protein) [32]. In addition, a set of Mn-regulated proteins were identified in the Mn-tolerant stylo genotype through proteomic analysis. These proteins are mainly involved in defense responses, photosynthesis, carbon fixation, metabolism, cell wall modulation, and signaling [35]. Further analysis shows that some of the identified proteins related to the phenylpropanoid pathway, including phenylalanine ammonia-lyase (PAL), chalcone synthase (CHS), chalcone–flavonone isomerase family protein (CFI), and isoflavone reductase (IFR), are regulated by external Mn in stylo [35]. As secondary metabolites, such as phenolics, flavonoids, phenylalanine, and callose, have been reported to be regulated by excess Mn in plants [12,16,98], the regulation of the phenylpropanoid pathway seems to facilitate plants' adaptations to Mn toxicity. Furthermore, combined with the physiological and proteomic analysis, the molecular responses involved in stylo adaptation to Mn toxicity are suggested to include enhancing defense responses and phenylpropanoid pathways, adjusting photosynthesis and metabolic processes, and modulating protein synthesis and turnover [35]. Despite the advances in the identification of various genes and proteins responding to Mn toxicity, there remains a scarcity of work designed to investigate how these genes are involved in plant tolerance to Mn toxicity, and future work is needed in these areas.

### **9. Future Perspectives**

Of the mineral nutrients essential for plant growth, Mn can cause phytotoxicity at excess levels, especially in acid soils. Even with the examination of the physiological and molecular mechanisms

and characterization of genes controlling Mn tolerance over the last few decades, relatively little is known about the molecular mechanisms regulating Mn homeostasis and detoxification in plants, which are critical to allow plants to adjust their Mn requirements and to avoid toxicity. Furthermore, as many genes responsible for Mn transport and distribution are not or are only slightly responsive to external Mn, future work is required to elucidate the possible regulatory mechanisms, such as transcriptional regulatory networks and post-translational protein modifications (e.g., phosphorylation, ubiquitination, and glycosylation), by which these components facilitate plant adaptions to changing Mn levels.

Although some genes have been implicated in Mn detoxification via ectopic expression in model yeast cells or Arabidopsis, the exact roles of these genes need to be determined at both the cellular and whole-plant levels, considering molecular and physiological aspects in planta. Aside from the model plants Arabidopsis and rice, candidate genes in other crop species should be identified to clarify their roles in Mn acquisition and detoxification, which might be more complicated depending on the physiological, biochemical, and molecular responses in different crops. Once identified, these genes can potentially be used to breed crop varieties with high Mn acquisition efficiency under Mn deficiency in alkaline soils, or with increased Mn tolerance under Mn toxicity in acid soils. Additionally, in some hyperaccumulator plants that can store high levels of toxic metals without displaying obvious toxicity, excess Mn has been shown to accumulate in the non-photosynthetic tissues for detoxification [99,100]. However, the mechanisms underlying Mn hyperaccumulation and the responses of hyperaccumulators to Mn remain poorly understood. Candidate genes responsible for Mn detoxification in Mn-hyperaccumulator plants have yet to be reported. These are some of the future directions that should be taken into account, as these resources can be exploited to develop genetically engineered plants used for Mn phytoremediation.

To date, most of the studies conducted to investigate gene functions in Mn detoxification have mainly focused on Mn transport, distribution, or homeostasis. Genes associated with other pathways, such as biological regulation and signal transduction, photosynthesis, carbohydrate and energy metabolism, and secondary metabolism, which can potentially influence Mn tolerance mechanisms, have received little attention. Future efforts to investigate these areas are of great importance for increasing our understanding of how plants detoxify Mn.

### **10. Conclusions**

Although Mn is an essential element for plants, excess Mn can cause phytotoxicity, inhibiting plant growth. This review shows that increasing plant Mn tolerance can be achieved by coordination of Mn absorption, translocation, and distribution, as well as by complex regulations of physiological changes and biochemical responses. This review highlights that Mn detoxification is regulated by a variety of genes and proteins associated with specific pathways, such as Mn transport and homeostasis, which can potentially be used to breed crop varieties with high Mn tolerance. This review also provides some of the future areas that could be taken into account in terms of gaining a better understanding of how plants tolerate Mn toxicity.

**Author Contributions:** Z.C. and G.L. designed the review; J.L., Y.J., R.D., and P.L. accessed the information; R.H., Z.W., and Z.C. revised the manuscript; J.L., Y.J., R.D., and X.L. designed the charts; J.L., Y.J., and Z.C. wrote the paper. All authors read and approved the final manuscript.**Funding:** This work was supported by the National Natural Science Foundation of China (31801951, 31861143013), the Young Talents Academic Innovation Project of Hainan Association for Science and Technology (QCXM201715), the Key Research and Development Program of Hainan (ZDYF2018048), the Central Public-interest Scientific Institution Basal Research Fund for CATAS (1630032017086), the Modern Agro-Industry Technology Research System (CARS-34), the Construction of World First Class Discipline of Hainan University (No.RZZX201905), and the Integrated Demonstration of Key Techniques for the Industrial Development of Featured Crops in Rocky Desertification Areas of Yunnan–Guangxi–Guizhou Provinces.

**Acknowledgments:** We sincerely thank American Journal Experts (www.aje.com) for their help with English language editing.

**Conflicts of Interest:** The authors declare no conflict of interest.

### **References**


© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Combined E**ff**ect of Cadmium and Lead on Durum Wheat**

### **Alessio Aprile 1, Erika Sabella 1,\*, Enrico Francia 2, Justyna Milc 2, Domenico Ronga 2, Nicola Pecchioni 2, Erika Ferrari 3, Andrea Luvisi 1, Marzia Vergine <sup>1</sup> and Luigi De Bellis <sup>1</sup>**


Received: 22 October 2019; Accepted: 22 November 2019; Published: 24 November 2019

**Abstract:** Cadmium (Cd) and lead (Pb) are two toxic heavy metals (HMs) whose presence in soil is generally low. However, industrial and agricultural activities in recent years have significantly raised their levels, causing progressive accumulations in plant edible tissues, and stimulating research in this field. Studies on toxic metals are commonly focused on a single metal, but toxic metals occur simultaneously. The understanding of the mechanisms of interaction between HMs during uptake is important to design agronomic or genetic strategies to limit contamination of crops. To study the single and combined effect of Cd and Pb on durum wheat, a hydroponic experiment was established to examine the accumulation of the two HMs. Moreover, the molecular mechanisms activated in the roots were investigated paying attention to transcription factors (bHLH family), heavy metal transporters and genes involved in the biosynthesis of metal chelators (nicotianamine and mugineic acid). Cd and Pb are accumulated following different molecular strategies by durum wheat plants, even if the two metals interact with each other influencing their respective uptake and translocation. Finally, we demonstrated that some genes (*bHLH 29, YSL2, ZIF1, ZIFL1, ZIFL2, NAS2* and *NAAT*) were induced in the durum wheat roots only in response to Cd.

**Keywords:** cadmium; lead; nicotianamine; mugineic acid; heavy metal; toxic metal; durum wheat

### **1. Introduction**

The distribution of heavy metals (HMs) in soils is variable from one place to another. In some regions, the HM natural background is higher than in other ones, but industrialization and human activities had strongly, in last decades, modified the concentration of many HMs worldwide [1]. A concern with HMs is that they can easily enter the food chain through consumption of vegetables and plant parts. Indeed, the toxicity is due mainly to chronic exposure by eating HM-contaminated foods. Moreover, since low levels of HMs in soil generally do not affect plant growth and development (no visible symptoms), HMs could endanger human health [2] if adequate counteractions are not implemented. Soil metal contamination usually occurs with a combination of different metals. Cadmium (Cd) and lead (Pb) are considered environmental hazards, as they are toxic for humans and other living organisms [3,4] and the Codex Alimentarius (CDX 193-1995, Amended 2019) has set a maximum level of 0.2 mg kg−<sup>1</sup> for both Cd and Pb in wheat [5].

Cd is an element of group II B in the periodic table and its atomic number is 48, while Pb belongs to group IV A and its atomic number is 82. Cd and Pb can form complexes with other compounds. In particular, Cd could form complexes with ammonia, amines, halide ions and cyanide [6]. In soils, Pb makes complexes with inorganic constituents (e.g., HCO3 <sup>−</sup>, CO3 <sup>2</sup><sup>−</sup>, SO4 <sup>2</sup><sup>−</sup> and Cl−), or may occur as organic ligands [7].

Although, Cd and Pb are not essential elements, plants are able to adsorb these metals from the soil and store them on different edible organs [8,9]. For these reasons, the control of HM accumulation in plant edible organs is a key point to preserve human health.

Plants respond to HM toxicity activating several physiological and molecular mechanisms. Such responses include immobilization, exclusion, chelation and compartmentalization of the metal ions, and the expression of common stress-related genes such as those involved in ethylene pathway and genes coding for stress proteins [10]. The current approach in risk-evaluation of HMs accumulation in plants is almost always based on the effects of single contaminants [11], but the combined effect of HMs is still low investigated.

Durum wheat is a staple food used to produce pasta (mainly in Europe and America), couscous and freekeh (Africa), bulgur (Asia) and bread (South Italy); minor crop if compared to bread wheat, it is mainly cultivated in Europe (Italy and France), America (Canada, USA, Mexico and Argentina), North Africa and Asia (Ukraine, Russia, Kazakhstan Indi and, China). These regions have different climate conditions and soil types with variable levels of toxic metals. In recent years, many authors have reported information about metal accumulation in durum wheat tissues, such as Cd [12,13], copper [14], arsenic [15], nickel [16], Pb, copper and chromium [17]. However, little is known about the combined effect of these metals on plant development, metal compartmentalization in plant tissues and molecular responses. Recently Shafiq et al. [18] reported how the expression of Heavy Metal ATPase 2 and ATP-Binding Cassette and promoter methylation could have a central role in Cd, Pb and zinc accumulation.

The aim of this work was to investigate the uptake and translocation of Cd and Pb, their interaction in root and leaf tissues and the molecular mechanisms activated by the aboveground presence of one or both metals during the growth of durum wheat plants. We employed two near-isogenic lines (NILs) [19] with an opposite behavior concerning Cd accumulation in leaves, low Cd (L-Cd NIL) and high Cd (H-Cd NIL), respectively. Since Cd accumulation is strongly affected by specific genomic regions [20], we should observe similar accumulation trends in these plants if the same genome regions are involved in Pb accumulation. Moreover, we added to the experiment design two commercial cultivars already characterized for their accumulation and responses to Cd [13] and, with this experiment, we evaluated how the contemporaneous presence of the Cd and Pb can affect each other.

### **2. Results**

### *2.1. Levels of Cd and Pb in Root and Leaf of Wheat Plants*

Samples were collected 42 days after germination, at the tillering stage, from plants grown in hydroponic solutions with the addition of Cd or Pb or with both metals at the concentrations of 0.5 and 2.0 μM, respectively. Figures 1 and 2 show the concentration of the two HMs in roots and leaves of the wheat plants; in roots, Cd and Pb concentrations range from 10 to 40 μg/g dry weight. The presence of a higher level of Cd and Pb in roots is evident (Figures 1 and 2): approximately, the concentration in roots is ten-times higher than in leaves. The Cd concentration in leaves expressed as μg/g dry weight is also twice as high as that of Pb and about four times if expressed as molarity because of the different atomic weight of the two metals.

L-Cd NIL, the near isogenic line characterized by the ability to accumulate a low level of Cd in leaves, collected, as expected, a low level of Cd in leaves compared to H-Cd NIL and to Svevo (Figure 1), whereas it showed the presence of a high level of Pb in leaves in comparison with all other genotypes (Figure 2), suggesting the presence of different molecular mechanism for the transport of the two metals. In L-Cd NIL the co-presence of Pb reduced the Cd accumulation in L-Cd NIL leaves (Figure 1).

Despite H-Cd NIL theoretically shares about the 95% of the genome with L-Cd NIL, it accumulates a high level of Cd in leaves [17], as confirmed in Figure 1. The co-presence of Pb did not alter significantly the concentration of Cd, which remained at comparable levels in both roots and leaves. Instead, L-Cd NIL and H-Cd NIL showed a similar behavior concerning the uptake and translocation of Pb and the co-presence of Cd altered considerably the Pb accumulation only in L-Cd NIL leaves (Figure 2).

The accumulation of Cd in Creso was influenced by the co-presence of Pb in hydroponic solution: when Creso was treated with both metals, the Cd concentrations in roots and leaves were slightly lower indicating a negative effect of Pb on Cd uptake/translocation.

**Figure 1.** Cadmium (Cd) concentration in roots and leaves of low cadmium near-isogenic line (L-Cd NIL), high cadmium near-isogenic line (H-Cd NIL), Creso and Svevo. Cd concentrations in durum wheat genotypes grown in standard hydroponic solution in the presence of Cd 0.5 μM, lead (Pb) 2.0 μM or in the presence of both heavy metals (HMs) (Cd 0.5 μM plus Pb 2.0 μM). Roots and leaves were collected 42 days after germination (at the tillering stage). Cd concentration was quantified by inductively coupled plasma mass spectrometer (ICP-MS). Statistical analysis was performed through ANOVA (*p*-value < 0.05, *n* = 3) followed by Tukey-HSD post hoc test. Different letters correspond to statistically different means.

**Figure 2.** Pb concentration in roots and leaves of L-Cd NIL, H-Cd NIL, Creso and Svevo. Pb concentrations in durum wheat genotypes grown in standard hydroponic solution in the presence of Cd 0.5 μM, Pb 2.0 μM or in the presence of both HMs (Cd 0.5 μM plus Pb 2.0 μM). Roots and leaves were collected 42 days after germination (at the tillering stage). Pb concentration was quantified by ICP-MS. Statistical analysis was performed through ANOVA (*p*-value < 0.05, *n* = 3) followed by Tukey-HSD post hoc test. Different letters correspond to statistically different means.

Svevo, compared to the other genotypes, resulted more sensitive to the occurrence at the same time of two metals in the hydroponic solution. Indeed, in Svevo the addition of a second metal strongly reduced the accumulation of the other one. This behavior was observed in roots (both Cd and Pb) and in leaves (only Cd was significantly reduced after the application of Pb).

An important parameter to study HMs uptake and translocation in plants is the translocation factor (Figure 3); it is the ratio of the metal concentration in other plant tissues in relation to roots [3]. Figure 3 indicates that the translocation factors from roots to leaves for Cd and Pb were clearly lower than 1.0 (ranging around 0.1 or less as it results from the values on the axes) indicating a robust limitation in Cd and Pb transport in durum wheat and an immobilization in the root cells; this strategy is widely used by plants to protect the photosynthetic tissues from damages caused by the HMs [21]. Anyway, translocation factor values showed that in the genotypes L-Cd NIL and Creso, Cd and Pb were translocated from root to shoot with a similar ratio; conversely, in Svevo and H-Cd NIL, Cd was translocated from root to shoot more efficiently than Pb compared with the other genotypes (Figure 3a). This is an expected result since Svevo and H-Cd NIL are well-known genotypes with high grain-Cd accumulation [13,17]. During the combined treatment with Cd and Pb, the genotype H-Cd NIL kept higher translocation factor values for Cd (Figure 3b), while in Svevo, the translocation factor of Pb became significantly higher than the translocation factors of Cd (Figure 3b). This is explainable

since in Svevo the HMs combined treatment affected Cd accumulation more at leaves level than in roots (Figure 1) causing a decrease in translocation factor values. In contrast, the combined treatment impacted Pb accumulation at root level more significantly than in leaves (Figure 2), determining an increase in translocation factors values.

Such data may represent a first evidence of the existence of different response mechanisms to Cd and Pb.

**Figure 3.** Translocation factor of Cd and Pb from root to shoot of wheat plants, (**a**) after the single treatments with Cd and Pb and (**b**) after the combined treatment with the two HMs. The significant differences, between the single heavy metal treatments, were highlighted according to Student's *t*-test (\* *p* < 0.05; \*\* *p* < 0.01; \*\*\* *p* < 0.001).

### *2.2. Gene Expression in Response to Cd and Pb*

The differences in Cd and Pb accumulation observed in the durum wheat genotypes may be induced by differential expression of gene categories involved in metal ion response and transport. The role of these genes has already been investigated in tissues of plants treated with Cd [22]. To study the molecular response to Cd, Pb and their combined effect, we considered the level of expression of genes already known to be involved in HMs responses. We focused the attention on transcription factors (bHLH and WRKY family), eight metal ion transporters and genes coding for the enzyme responsible of the synthesis of nicotianamine and mugineic acid, that are two typical metal chelators of graminaceous plants [23]. The expression levels of these genes were analyzed both in roots and leaves, but we observed no expression or no differential expression in durum wheat leaves, suggesting a tissue-specific regulation/expression. Below, only the root transcription data were reported.

### 2.2.1. Expression of the Transcription Factors Basic Helix-Loop-Helix (bHLH) and WRKY33

As shown in Figure 4, the expression of *bHLH29*/*FIT* and *bHLH38*/*ORG2* was clearly up-regulated in the roots of the four analyzed genotypes when treated with Cd and with Cd plus Pb; on the contrary, *bHLH47*/*PYE* (Figure 4) was only up-regulated in the genotype H-Cd NIL both after a treatment with Cd or Pb and following the combined stress determined by Cd plus Pb. *WRKY33*, member of the WRKY transcription factors family, did not show a significant modulation in roots of the four genotypes during treatments with the HMs (Figure 4).

**Figure 4.** Relative expression as fold change (FC) of the *bHLH29*/*FIT*, *bHLH38*/*ORG2*, *bHLH47*/*PYE* and *WRKY33* genes in root tissues of the durum wheat genotypes (L-Cd NIL, H-Cd NIL, Creso and Svevo) grown in the presence of Cd 0.5 μM, Pb 2.0 μM or in the presence of both HMs (Cd 0.5 μM plus Pb 2.0 μM). Error bars indicate standard deviation of the mean of three technical replicates resulting from a bulk of three biological replicates. ANOVA results were reported basing on their statistical significance. \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001.

### 2.2.2. Expression of HMs Transporters

Plants treated with HMs showed no significant change in transcripts of the genes coding for the HM transporters ZIP4 and ZTP29 in root tissues (Figure 5). *YSL1* was slightly up-regulated in the roots of the L-Cd NIL and Creso when treated with Cd or with Cd plus Pb (Figure 5); conversely, expression level of *YSL2* was strongly up-regulated in roots of L-Cd NIL and Creso when treated with Cd. The presence of both Cd and Pb resulted in a significant increase of the transcript levels in

roots of L-Cd NIL; in roots of the cultivar Creso, the combined stress induced a slight upregulation (Figure 5). A slight up-regulation was observed also in H-Cd NIL and Svevo roots treated with Cd and in the combined treatment Cd plus Pb (Figure 5). The vacuolar zinc transporter genes *ZIF* and *ZIF*-like genes (*ZIFL1* and *ZIFL2*) were strongly up-regulated in the roots of the four durum wheat genotypes treated with Cd and Cd plus Pb (Figure 6). The transcript levels were significantly higher in L-Cd NIL and Creso (Figure 6). Finally, the plasma membrane-localized transporter *HMA5* was significantly up-regulated in roots when treated both with one HM, or with the combined Cd plus Pb (Figure 6).

**Figure 5.** Relative expression (FC) of the *ZIP4*, *ZTP29*, *YSL1* and *YSL2* genes in root tissues of the durum wheat genotypes (L-Cd NIL, H-Cd NIL, Creso and Svevo) grown in the presence of Cd 0.5 μM, Pb 2.0 μM or in the presence of both HMs (Cd 0.5 μM plus Pb 2.0 μM). Error bars indicate standard deviation of the mean of three technical replicates derived from a bulk of three biological replicates. ANOVA results were reported basing on their statistical significance. \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001.

**Figure 6.** Relative expression (FC) of the *ZIF1*, *ZIFL1*, *ZIFL2* and *HMA5* genes in root tissues of the durum wheat genotypes (L-Cd NIL, H-Cd NIL, Creso and Svevo) grown in the presence of Cd 0.5 μM, Pb 2.0 μM or in the presence of both HMs (Cd 0.5 μM plus Pb 2.0 μM). Error bars indicate standard deviation of the mean of three technical replicates derived from a bulk of three biological replicates. ANOVA results were reported basing on their statistical significance. \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001.

2.2.3. Expression of the Nicotianamine Synthase Genes (NAS) and Nicotianamine Aminotransferase (NAAT)

In the current study, the transcripts levels of the nicotianamine synthase genes (*NAS2*, *NAS3* and *NAS4*) confirmed their up-regulation in roots of plants treated with Cd (Figure 7) as observed by other authors [19]. Interestingly, a significant increase of the transcript levels for the genes *NAS2*, *NAS3* and *NAS4* were also found in roots of plants treated with Cd plus Pb (Figure 7). Moreover, the gene *NAS3* resulted modulated in response to Pb treatment too, with a minor induction in roots (Figure 7). The transamination of nicotianamine produces mugineic acid and the enzyme that catalyzes the synthesis is called nicotianamine aminotransferase (NAAT); the relative gene is expressed at highest levels in roots of L-Cd NIL and Creso grown in the presence of Cd and Cd plus Pb; a significant induction was also found in H-Cd NIL and Svevo exposed to Cd and Cd plus Pb (Figure 7).

**Figure 7.** Relative expression (FC) of the nicotianamine synthase genes (*NAS2*, *NAS3*, *NAS4*) and nicotianamine aminotransferase gene (*NAAT*) in root tissues of the durum wheat genotypes (L-Cd, H-Cd, Creso and Svevo) grown in the presence of Cd 0.5 μM, Pb 2.0 μM or in the presence of both HMs (Cd 0.5 μM plus Pb 2.0 μM). Error bars indicate standard deviation of the mean of three technical replicates derived from a bulk of three biological replicates. ANOVA results were reported basing on their statistical significance. \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001.

### **3. Discussion**

The data support the hypothesis that the effects produced by combinations of HMs could have a different impact on both accumulation and gene expression in comparison to the individual effects of each metal. The analyzed durum wheat genotypes showed different distribution of Cd and Pb in roots and leaves when treated with the two HMs in comparison with a treatment with only one metal. In general, the four genotypes showed a reduced accumulation of both Cd and Pb in the combined treatment (Figures 1 and 2). Several studies described that the presence of one metal influenced the uptake of another metal [24,25]. Xie et al. [25] investigated the effects of combined HMs toxicity on two rice genotypes differing in Cd accumulation; they found that the application of Pb, Cd, chromium (Cr) and copper (Cu) significantly affected grain Cd accumulations. In rice, Zeng et al. [26] also reported a significant Cd and Pb reduction in grains when exposed to both metals. Other studies were carried out on cucumber (*Cucumis sativus* that could retain greater amount of metals in the roots due to its root morphology) [27] to assess HMs toxicity in soils contaminated by Cu, Cd and Pb separately and in combinations; according to their results, bioaccumulation of one metal was influenced by the presence of other metals and, in general, the HMs accumulation patterns reflected antagonistic and/or synergistic plant's responses. In the binary combination of Cd and Pb, they found a synergistic response with a reduction of toxicity effect [28]. Cd/Pb synergisms have been previously reported by Zaray et al. [29]. On *Lemna minor*, a metal pollution sensitive plant, the combined toxicity of Pb and Cd was found to be less effective when compared to the toxicity of the individual treatment [30].

The data about Cd and Pb accumulation in leaves highlighted different behaviors among genotypes in relation to the two toxic metals. Svevo and H-Cd NIL are good accumulators of both Cd and Pb in leaves. On the contrary Creso accumulates lower level of Cd and Pb if compared to the other genotypes. The L-Cd NIL has a contrasting behavior: accumulates low level of Cd and high level of Pb. It is interesting to note that the expression levels of the tested genes in Svevo and H-Cd NIL are generally less up-regulated or not-regulated at all, suggesting that these genes are involved in some molecular mechanisms to stuck Cd and Pb at root level (e.g., genes involved in vacuole compartmentalization of toxic metals).

These differences (both in accumulation and distribution of HMs) could be, in part, due to differential expression of genes involved in HM uptake, cellular sequestration and translocation from root to shoot. Since among the HMs, Cd is accumulated in the grain of durum wheat to levels exceeding the Codex Alimentarius Commission standards [5] Cd uptake, cellular sequestration and translocation have been thoroughly studied [31,32]; some of the genes, with a key role in these physiological steps in durum wheat plants during Cd treatment [22], were investigated during the combined exposure to Cd and Pb.

Cd can enter root cells as Cd2<sup>+</sup> through ZIP (zinc regulated transporter/iron regulated transporter-like protein) transporters or as Cd-chelates through YSL (Yellow-Stripe 1-Like) proteins [33]. Some investigations have proven that the ZIP family transporters participate in Cd absorption and accumulation in plants [34]; in our work, the selected gene *ZIP4* was not significantly regulated (Figure 5) in response to Cd treatment. Yamaguchi et al. [35] also found that genes coding for ZIP transporters did not change their expression patterns in *Solanum torvum* roots during treatment with low Cd concentrations and they postulated that the absence of changes in these metal transporters may explain why the mild Cd exposure did not induce serious competitive inhibitory effects on metal ion homeostasis in roots. The same effect due to the exposure to a low HM concentration could explain the unchanged expression of the gene *ZTP29* (Figure 5). This gene, coding for a zinc transporter with homology to the *Arabidopsis ZTP29*, is localized into the endoplasmic reticulum and it is thought to play a role in the unfolded protein response [36], Liu et al. [37] found that Cd up-regulated *ZTP29* in roots of *Cosmos bipinnatus* Cav. When they are under 40 μmol/L Cd stress (80 times more concentrated than the concentration used in this work).

Instead, the genes coding for the YSL (Yellow-Stripe 1-Like) proteins responsible for the transport of the Cd-chelates resulted up-regulated in Cd and Cd plus Pb treatments with higher FC values in the genotypes L-Cd NIL and Creso in which also *YSL2* was vigorously activated (Figure 5). This upregulation in response to Cd treatment is in accordance with RNA sequencing data reported in Aprile et al. [22]. The YSL family of transporters represents a candidate for the transport of nicotianamine (NA)–metal chelates across plant cell membranes [38]; several members of the YSL family are localized to the plasma membrane and function as transporters of metals that are bound to the metal chelator nicotianamine or the related set of mugineic acid family chelators known as phytosiderophores [39]. Other YSL members are localized to the vacuole membranes and to the internal membranes and may play a role in detoxification by HM sequestration in the vacuole [40]; this compartmentalization mechanism could contribute with the characteristic trait of low Cd-accumulation in durum wheat grain [13]. The two genotypes (L-Cd NIL and Creso) with lower Cd and Pb accumulation in leaves, had also the higher level of expression of the gene *YSL2*, suggesting a possible regulatory role in Cd and Pb compartmentalization in roots. The treatment with Pb did not affect the expression level of the *YSL* transporters while the Cd-Pb combined treatment affects the *YSL2* expression in the L-Cd NIL and Creso by reducing the amount of mRNA in comparison with the expression level in the single Cd treatment; the exposure to the combined HMs can activate simultaneously several nonspecific defense systems and it is reported that the interactions among HMs affect their uptake and accumulation in plants [25,26,31,41]. Other NA vacuolar transporter genes are the *ZIF* and *ZIF*-like genes, their expression levels were up-regulated by Cd and Cd plus Pb treatments in all the analyzed genotypes while the Pb treatment did not change expression pattern (Figure 6); so it is reasonable to assume that the Cd-nicotinamine chelates could enter into the vacuoles through the ZIF and ZIFL.

HMs are loaded from the symplast into the xylem by heavy metal P1B-ATPases, known as heavy metal ATPases (HMAs) that play an important role in metal transport in plants [20,33].

In our experiment, the gene coding for the transporter HMA5 was upregulated both in Cd and Pb single treatment and the combined treatment increased the amount of mRNA if compared with the HM single treatment as if there was a cumulative effect (Figure 6). Functional studies on the HMAs have shown that these transporters can be divided into two subgroups based on their metal-substrate specificity: a copper (Cu)/silver (Ag) group and a zinc (Zn)/cobalt (Co)/Cd/Pb group [42]; this indication is in accordance with our data since *HMA5* was induced both by Cd and Pb. Besides genes coding for HMs transporters, previous studies have characterized several transcription factors (TFs) involved in Cd response: ERF, WRKY and bHLH TF families [43]. Obtained gene expression patterns supported the involvement of these genes in the response to Cd stress since *bHLH29*/*FIT*, *bHLH38*/*ORG2* and *bHLH47*/*PYE* were significantly induced only in response to Cd treatment (Figure 4). A significant up-regulation in response to Pb treatment was recorded exclusively in the genotype Svevo for the gene *bHLH38*/*ORG2* (Figure 4). For the TF *WRKY33* not significant induction was observed (Figure 4). Since many transcription factors are transiently regulated by stresses/treatments, the long-term exposure to Cd and Pb and the sample collection at 42 days after germination were not suitable to observe a transcription variation. In *Arabidopsis thaliana* treated with Cd, real-time PCR analyses demonstrated that the *WRKY13* transcript was rapidly induced by Cd stress, reaching the maximum level after 1 h of Cd treatment and gradually decreasing, thereafter [44]. On the other hand, Long et al. [45] described how the co-overexpression of *FIT* and *ORG2* enhanced the expression of nicotianamine synthase 1 (*NAS1*) and *NAS2*, resulting in the accumulation of nicotianamine, a crucial chelator for Fe transportation and homeostasis.

Figure 7 showed a strong induction for the genes *NAS2*, *NAS3* and *NAS4* in response to Cd treatment while no gene expression modulation was recorded in response to Pb treatment. These data suggested that the molecular mechanisms regulated by the bHLH TFs are well conserved between *Arabidopsis* and durum wheat and also gave evidence of a detoxification mechanism specific for Cd stress since Pb treatment did not activate the synthesis of nicotianamine. According to Pal and Ray [46], phytochelatins synthesis is influenced by the metal ion treatment and a number of phytochelatins variants have been found among plant species. Nicotianamine (NA) is a non-proteogenic amino acid chelator having more than one binding centers, which confer high affinity for Fe, but also for

other metals such as Zn, Cu, Mn, Ni and Cd [46]. Another chelator, involved in iron uptake in graminaceous plant species, is the mugineic acid [47]. In the mucigenic acid biosynthetic pathway, nicotianamine aminotransferase (NAAT) is implicated in the formation of 2 -deoxymugineic acid (DMA) from nicotianamine [48]. With our expression data, we described that *NAAT* was highly expressed in root tissues of all the analyzed genotypes treated with Cd (Figure 7) indicating a critical role for the mucigenic acid in response to Cd.

### **4. Materials and Methods**

### *4.1. Genetic Materials*

To identify the effects of the combined stress of Cd and Pb in durum wheat (*Triticum turgidum* L. subsp. *Durum*), a pot experiment was conducted in a growth chamber. Two low grain Cd accumulation (L-Cd NIL and Creso) and two high grain Cd accumulation (H-Cd NIL and Svevo) were analyzed. Creso and Svevo accession numbers are, respectively, K-53049 and RICP-01C0107074, and all pedigree information is browsable at CIMMYT database (http://www.wheatpedigree.net). L-Cd NIL and H-Cd NIL are two near-isogenic lines (NILs) of durum wheat (*Triticum turgidum* L. subsp. *durum*) that differ in grain Cd accumulation TL 8982-H (H-Cd NIL) and TL 8982\_L (L-Cd NIL) [17].

### *4.2. Experimental Design*

Cd and Pb management to the four genotypes was set up by hydroponic system. After external sterilization, seeds were germinated in Petri dishes with moist filter paper, in the dark at 8 ◦C. After germination (6–7 days), plantlets were located into 0.4 L plastic pots (7 cm × 7 cm × 8 cm) filled with perlite, moistened with tap water, and immediately transferred to the 10 L polyethylene tanks of the hydroponic system, as described by Harris and Taylor [49] with little modifications. In each pot three seedlings were lodged in and for each treatment, three different pots were considered for three biological replicates. The positions of the pots in the growth chamber were completely randomized and changed weekly with a new randomization. Plants were grown in the growth chamber under long days, 16 h light/8 h night, 21 ◦C/16 ◦C. The hydroponic solution was given with systematic pauses, irrigating for 15 min every 2 h during the all day; while during the night no fertigation. In this way, the perlite substrate was constantly dampened with hydroponic solution, avoiding stagnation. The nutrient solution was prepared using reverse osmosis (RO) water (<30 μS cm−1) and contained: 0.3 mM NH4NO3, 0.25 mM KNO3, 0.1 mM K2SO4, 50 μM KCl, 1.0 mM Ca(NO3)2, 0.3 mM Mg(NO3)2, 100 μM Fe(NO3)3, 1.0 μM MnSO4, 10.0 μM H3BO3, 10.0 μM ZnSO4, 0.2 μM Na2MoO4, 2.0 μM CuSO4, 2.0 μM Cu(NO3)2, 0.1 mM K2HPO4, 138.6 μM *N*-(2-hydroxyethyl) ethylenediaminetriacetic acid (HEDTA), and 2 mM 2-(*N*-Morpholino) ethanesulfonic acid hydrate. After preparation of solution, pH was adjusted by 1.42 mM KOH.

The pH of the nutrient solution and its electrical conductivity (EC) were constantly monitored every 2 days, EC was used to estimate water depletion by keeping EC in the main tank between 550 and 600 μS cm<sup>−</sup>1, and nitric acid (1% *v*/*v*) was used to adjust pH between 5.5 and 6.0, when needed.

In cultivated fields the presence of Cd and Pb is usually not toxic for the crops. In this experiment plants were treated by adding to the nutrient solution 0.5 μM CdCl2, or 2.0 μM Pb(NO3)2, or even both in the case of the double metal stress. These concentrations do not cause a significant toxic effect on root and leaf biomass as reported respectively by Harris and Taylor [49] and Sun et al. [50], while Cd 0.5 μM has been employed in previous works of the University of Salento [12,22].

The control plants were cultivated without Cd or Pb in the same hydroponic solution. Hydroponic solution was constantly aerated. One plant for each pot (three for each treatment) was sampled 42 days after germination, at tillering stage (roots and leaves). Roots were easily extracted from perlite substrate and washed manually to remove the perlite beads adherent to roots, and possible excess of hydroponic solution. Leaf samples were washed immediately on harvest in RO water for 30 s, while root samples were triple rinsed (RO water, 1 min; 1 mM CaCl2, 5 min; RO water, 1 min), and blotted dry. Samples for quantitative RT-PCR were frozen in liquid nitrogen and then stored at −80 ◦C.

### *4.3. Inductively Coupled Plasma Mass Spectrometry (ICP-MS) Analysis*

Measurements of Cd and Pb uptake in roots were performed by using X SeriesII inductively coupled plasma mass spectrometer (ICP-MS; Thermo Fisher Scientific, Waltham, MA, USA) equipped with Peltier cooled (3 ◦C) spray chamber. Samples were introduced by the autosampler CETAC ASX 520 into the nebulizer, and the positively charged ions were then produced by a high-temperature, inductively coupled plasma. The ions passed through a sampling cone interface into a high-performance quadrupole mass spectrometer that is computer-controlled to carry out multi-element analysis. Data were analyzed by PlasmaLab software. The instrument was tuned daily with an ICP-MS tuning solution. Yttrium in HNO3 4% (100 ppb) was used as internal standard. Cd and Pb standards ranging from 0.2 to 100 ppb were freshly prepared before each analysis and used to build calibration curve. Each sample was analyzed at least in three independent measurements and each experiment comprised three repetitions. Results are given as mean value ± standard deviation.

Each sample was mineralized by a microwave-assisted procedure performed with an ultraWAWE microwawe digestion system (Milestone Inc., Shelton, CT, USA), as recently reported by Durante et al. [51]. The sample (~300 mg) was accurately weighed in the microwave-quartz vessels before adding 2 mL of ultrapure HNO3 65% *w*/*w* and 4 mL of Milli-Q water. At the end of the digestion process, an almost colorless, pale yellow sample was obtained. The solution was then diluted up to 10 g with Milli-Q water in test tubes. Between each mineralization cycle, a washing cycle was carried out.

Ultrapure HNO3 65% *w*/*w* was obtained from analytical grade nitric acid (Carlo Erba, Milan, Italy) after sub-boiling distillation performed with a sub-boiler SAVILLEX DST 1000 (Savillex Corp. Eden Prairie, MN, USA) apparatus.

### *4.4. Total RNA Isolation, cDNA Synthesis and qPCR Analysis of Gene Expression*

To evaluate the response of durum wheat plants to Cd and Pb treatments, we carried out a transcriptomic analysis in a small, but well-defined, group of genes [22]. Total RNA was extracted from root and leaf tissues using TRIZOL reagent according to the method published by Marè et al. [52]. To assess RNA quality and quantity, several dilutions of each sample were analyzed using the Agilent RNA 6000 nano Kit and Agilent Bioanalyzer 2100. cDNA synthesis was performed using TaqMan® Reverse Transcription Reagents (Applied Biosystems, Foster City, CA, USA). qPCR was performed with the Power SYBR Green RT-PCR Master mix (Applied Biosystems, Foster City, CA, USA) according to the manufacturer's instructions. To calculate the relative expression levels between a reference sample and the related treatments, the fold change (FC) formula was used:

$$\mathsf{RC} = \mathcal{Z}^{\ast - \Delta \Lambda \text{CT}}\_{\prime}$$

where ΔΔCT = (CTtargetgene − CTreferencegene) treatedsample − (CTtargetgene − CTreferencegene)controlsample.

Sequences related to genes coding for transcription factors (*bHLH29*, *bHLH38*, *bHLH47* and *WRKY33*), membrane transporters (*HMA5*, *YSL1*, *YSL2*, *ZIF1*, *ZIFL1*, *ZIFL2*, *ZIP4* and *ZTP29*) and genes involved in the metal chelator pathway (*NAS2*, *NAS3*, *NAS4* and *NAAT*) were downloaded from the site https://www.ebi.ac.uk/arrayexpress, at European Bioinformatics Institute (EMBL-EBI). The accession code is E-MTAB-7266. Then, the sequences were compared to the NCBI database using the BLAST algorithm and the most similar sequences were used to design the relative real-time PCR primers (Primer Express™ Software v3.0, Applied Biosystems, Foster City, CA, USA) Supplementary Table S1. TheNADH ubiquinone reductase gene was used as the reference gene to normalize the expression levels of the target genes.

### *4.5. Statistical Analyses*

Means of quantitative data related to Cd and Pb concentrations were determined for each tissue (root and leaf) and were subjected to two-way ANOVA analysis (genotype X treatment), followed by Tukey-HSD (honestly significant difference) post hoc test (*p* < 0.05). Translocation factor data were subjected to two-way ANOVA (genotype X treatment). A *t*-test was employed to find statistical differences between Cd and Pb translocation factors on each genotype. A one-way ANOVA and *t*-test were applied to expression gene data.

Analyses were achieved using R version 3.5.3.

### **5. Conclusions**

The mechanisms activated by plants to tolerate the presence of HMs were studied by exposing plants to not only a single HM but to a combination of Cd and Pb. Uptake and translocation strategies are not regulated by the same genes as suggested by the strong differences observed among genotypes in response to the two toxic HMs. In fact, the level of Cd in durum wheat roots and leaves is influenced by the co-presence of Pb and vice versa even if the phenomenon has different extent among genotypes.

Furthermore, nicotianamine and mucigenic acid seem to play a key role in response to Cd stress, probably by chelating the metal and avoiding its translocation to the plant shoots. The combined stress with Cd and Pb did not affect this mechanism, which appeared to be specific for Cd.

**Supplementary Materials:** Supplementary Materials can be found at http://www.mdpi.com/1422-0067/20/23/ 5891/s1.

**Author Contributions:** A.A., E.S., N.P. and L.D.B. have planned the experimental design. E.F. (Enrico Francia), J.M. and D.R. have grown plants in hydroponic conditions and collected samples for further investigations. E.F. (Erika Ferrari) has carried out the chemical analysis on Cd and Pb concentration in tissues. M.V. and A.L. processed samples for mRNA extraction. E.S. and A.A. have run the real-time PCR expression analysis and the relative data analysis. A.A., E.S. and L.D.B. wrote the manuscript and N.P., E.F. (Enrico Francia), A.A. and L.D.B. reviewed and edited the final version.

**Funding:** This work was economically supported by MIUR (Italian Minister of University and Research) project "Sviluppo tecnologico e innovazione per la sostenibilità e competitività della cerealicoltura meridionale MIUR-UE (PON01\_01145/1-ISCOCEM)" and by Regione Puglia, "FutureInResearch" project "Frumento duro "Cappelli": valorizzazione delle componenti genetiche alla base della tolleranza allo stress idrico", code 2I19HY5. These funding bodies had no role in the design of this study, during its execution, analyses, interpretation of the data, or decision to submit results.

**Acknowledgments:** We thank Curtis Pozniak and John Clarke for the kind sharing of the near isogenic lines TL 8982-H (H-Cd NIL) and TL 8982-L (L-Cd NIL). We are thankful to 'Centro Interdipartimentale Grandi Strumenti (CIGS)' of the University of Modena and Reggio Emilia that supplied ICP-MS, and to Prof. Andrea Marchetti for his precious advice and FKV autoclave.

**Conflicts of Interest:** The authors declare no conflict of interest.

### **References**


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