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Sulfur assimilation and glutathione metabolism under cadmium stress in yeast, protists and plants

David Mendoza-Cózatl, Herminia Loza-Tavera, Andrea Hernández-Navarro, Rafael Moreno-Sánchez
DOI: http://dx.doi.org/10.1016/j.femsre.2004.09.004 653-671 First published online: 1 September 2005


Glutathione (γ-glu-cys-gly; GSH) is usually present at high concentrations in most living cells, being the major reservoir of non-protein reduced sulfur. Because of its unique redox and nucleophilic properties, GSH serves in bio-reductive reactions as an important line of defense against reactive oxygen species, xenobiotics and heavy metals. GSH is synthesized from its constituent amino acids by two ATP-dependent reactions catalyzed by γ-glutamylcysteine synthetase and glutathione synthetase. In yeast, these enzymes are found in the cytosol, whereas in plants they are located in the cytosol and chloroplast. In protists, their location is not well established. In turn, the sulfur assimilation pathway, which leads to cysteine biosynthesis, involves high and low affinity sulfate transporters, and the enzymes ATP sulfurylase, APS kinase, PAPS reductase or APS reductase, sulfite reductase, serine acetyl transferase, O-acetylserine/O-acetylhomoserine sulfhydrylase and, in some organisms, also cystathionine β-synthase and cystathionine γ-lyase. The biochemical and genetic regulation of these pathways is affected by oxidative stress, sulfur deficiency and heavy metal exposure. Cells cope with heavy metal stress using different mechanisms, such as complexation and compartmentation. One of these mechanisms in some yeast, plants and protists is the enhanced synthesis of the heavy metal-chelating molecules GSH and phytochelatins, which are formed from GSH by phytochelatin synthase (PCS) in a heavy metal-dependent reaction; Cd2+ is the most potent activator of PCS. In this work, we review the biochemical and genetic mechanisms involved in the regulation of sulfate assimilation-reduction and GSH metabolism when yeast, plants and protists are challenged by Cd2+.

  • Cadmium resistance
  • Cadmium accumulation
  • Sulfur assimilation pathway
  • Phytochelatins
  • Phytochelatin synthase
  • γ-Glutamylcysteine synthetase
  • Glutathione synthetase

1 Introduction

Some heavy metals such as Cu2+, Co2+, Fe3+, Mn2+ and Zn2+ are essential in trace amounts for cell metabolism, acting either as enzyme cofactors, mediating redox reactions, and interacting with nucleic acids and proteins [13]. Others, such as Cd2+, Pb2+, Hg2+ and Ag+, although lacking biological function, enter into the cell through the same transport systems used by essential heavy metals, altering cellular functions [46]. A commonly used definition of “heavy metal” is that referring to all chemical elements with a density greater than 5 g ml−1. Metals may also be classified in three types, depending on their reactivity with the functional groups of biomolecules. Class A metals (Al3+, Ca2+, Sr2+, Ba2+, La3+) show more reactivity with oxygen (O > N > S); class B metals (Cu1+, Hg2+, Ag1+) prefer sulfur (S > N > O); and class C metals (Fe3+, Ni2+, Zn2+, Cd2+, Cu2+) have an intermediate affinity [7].

The toxic effects of heavy metals depend on the time and concentration to which organisms are exposed. Most of the effects are related to their interaction with carboxyl and thiol groups of proteins, to their ionophoretic properties and to their ability to direct or indirectly generate free radicals and hence induce oxidative stress [79]. Organisms possess diverse mechanisms to maintain free metal concentrations at levels that do not exceed cellular requirements. These mechanisms include (a) cellular wall binding, (b) changes in ion permeability, (c) active extrusion, (d) bio-transformation, (e) extra- and intracellular chelation, and (f) compartmentation [1, 1014b].

One of the best described mechanisms against heavy metals toxicity in some yeast [1517], algae [18], photosynthetic protists [19], and plants [12, 14a, 14b, 20] involves their intracellular chelation by either GSH or phytochelatins (PCs), low molecular weight sulfur-containing peptides derived from GSH [12, 14a, 15, 21a, 21b], or both. These peptides may bind a variety of metals in the cytosol and, depending on the organism, the metal–PC or metal–(GSH)2 complexes are actively transported into the vacuole [21a–23]. Like metallothioneins, the PC–metal complexes may activate metal-requiring apo-enzymes; for this reason they have been associated with the regulation of intracellular levels of essential heavy metal and with detoxification of non-essential ones [24, 25].

GSH is present in all organisms participating in multiple metabolic processes; for example, intracellular redox state regulation, inactivation of reactive oxygen species (ROS), transport of GSH-conjugated amino acids and other molecules, and storage of sulfur and cysteine affording up to 90% of the non-proteic sulfur in the cell [26, 27]. GSH synthesis, starting from inorganic sulfate, requires the sulfur assimilation (SAP) and the cysteine biosynthetic (Cys) pathways (Figs. 1 and 2). The biochemical and genetic regulation of these pathways is complex and is affected by different stress situations such as heavy metal exposure, oxidative stress and sulfur or nitrogen deficiency [2831]. Several reviews have appeared which analyze the biochemical characteristics of the enzymes and the regulation of the genes involved in the SAP and Cys synthesis in yeast and plants [3238]. However, none of them have focused on the regulation at the enzymatic and genetic level of cysteine and glutathione synthesis under heavy metal exposure. Therefore, in the present work, we analyze and discuss recent advances in the knowledge of reactions, of enzyme characteristics and properties, and of the biochemical and genetic regulatory mechanisms involved in coping with heavy metal toxicity, specifically with Cd2+. Where appropriate, the lack of information and research on a particular subject is addressed. Biotechnological relevance of this knowledge resides in the possibility of developing organisms with high capacity of Cd2+ accumulation for bioremediation purposes.


Sulfate assimilation pathway (SAP) and biosynthesis of cysteine. Numbered reactions are described in the text.


GSH, PCs synthesis and ROS processing through the ascorbate–glutathione system. AA, amino acid; X, xenobiotic or compound attached to GSH by GSH-S-transferases. Numbered reactions are described in the text.

2 Sulfur assimilation

2.1 Sulfate uptake

Sulfate is co-transported into the cells with 3H+, in an energy-dependent process catalyzed by specific plasma membrane permeases (Fig. 1, reaction 1) [35, 39]. High (Km < 10 μM) and low (Km > 100 μM) affinity sulfate transporters (HAST and LAST) have been described in different organisms. Although in Saccharomyces cerevisiae both activities have been detected (Table 1) [40], only genes encoding HASTs (SUL1 and SUL2) and a third gene (SUL3) involved in the transcriptional regulation of SUL2 [41] have been found. Another gene very similar to SUL1 and SUL2 is found in the S. cerevisiae genome, but its function is still unknown [42].

View this table:

Kinetic parameters for some of the enzymes involved in yeast sulfate uptake and reduction, and Cys biosynthesis

EnzymeAffinity for different ligands (Ks, Km or Ki)V mOrganismReference
LAST, low affinity sulfate transporter350 μM7.5 nmol min−1 (dry weight)−1S. cerevisiae[40]
HAST, high affinity sulfate transporter4 μM7 nmol min−1 (dry weight)−1S. cerevisiae[40]
ATPS, ATP sulfurylase0.17 mM Embedded Image2.1 μmol min−1 (mg protein)−1S. cerevisiae[195]
0.07 mM (ATP-Mg)
1 μM (Ki, APS)
APSK, APS kinasen.a.68 nmol min−1 (mg protein)−1S. cerevisiae[196]
PAPSR, PAPS reductase19 μM (PAPS)7 nmol min−1 (mg protein)−1S. cerevisiae[197]
SiR, sulfite reductase30 μM1 nmol min−1 (mg protein)−1S. cerevisiae[80]
OAS TL, O-acetyl(homo)serinethiol lyasen.a.24.8 nmol min−1 (mg protein)−1S. cerevisiae[127]
CT-βS, CT β-synthase2.25 mM (hCys)1.2 μmol min−1 (mg protein)−1S. cerevisiae[198]
2.1 mM (serine)
CT-γL, CT γ-lyase0.25 mM (CT)420 nmol min−1 (mg protein−1)S. cerevisiae[199]
  • The enzymes with the most complete kinetic analysis are shown. n.a, non available.

In plants, after its uptake by roots, sulfate is distributed into different organs and, to be assimilated, it has to be reduced in a process performed in chloroplasts [36, 37]. In these organisms, sulfate transporters are encoded by multigene families, which proteins have not only different sulfate affinity, but also different localization, expression patterns, and regulation [38]. In Arabidopsis, HASTs, which are mainly involved in sulfate transport from the environment to root tissues, are encoded by the Sultr1 family, in which three members have been identified [4345]. LASTs, expressed in roots and leaves, seem responsible for the sulfate uptake from the apoplast to different tissues inside the plant and are encoded by the Sultr2 family, with two members [43]. The gene expression of sulfate transporters in plants is also regulated by the availability of sulfate in the external medium, by the sulfate intracellular requirements [41, 46] and by GSH [47, 48].

In the green algae Chlamydomonas reinhardtii two sulfate transporters have been identified, one in cells growing in sulfur sufficient medium and another in sulfur deficient medium [49a]. Analysis of their kinetic parameters suggests that a LAST is present in cells grown in a complete medium whereas a HAST is induced under sulfur starvation. In the protist Euglena gracilis, some activities of the sulfur activating system have been detected in mitochondria [50], but there is no information about the nature of the sulfate transporters in this organelle or in plastids.

Biochemical and genetic characterization of the sulfate transporters of chloroplast membranes has not been addressed either in algae or plants. This is unfortunate since the chloroplast is the major site of sulfate reductive assimilation in these organisms. Several lines of evidence suggest that the sulfate transport system in chloroplasts has different genetic origins and has undergone several changes in the course of the evolution of algae and lower and higher plants. In some algae, such as Chlorella vulgaris, Mosostigma viridae, Nephroselmis olivacea, and Prototheca wickerhamii, as well as the primitive liverwort Marchantia polymorpha, genes encoding proteins similar to CysT have been identified in the chloroplast genome. CysT is a component of the sulfate permease system in the cyanobacteria Synechococcus sp. strain PCC 7942 and Synechocystis sp. PCC6803. In M. polymorpha also the CysA gene has been identified. However, the other components, essential for the functioning of the bacterial sulfate permease system, CysW and the sulfate binding protein (sbp) have not been found in the chloroplast or nuclear genomes of these organisms (see references cited in [51]). In the unicellular red alga Cyanidioschizon merolae the genes cysW and cysT have been found in the chloroplast genome and the genes cysA and sbp have been detected in the nuclear genome [52]. In higher plants such as Arabidopsis none of the genes encoding a bacterial sulfate transport system have been identified. A nuclear-encoded gene Sultr4;1 has been suggested to code for a sulfate transporter in the chloroplast [53], localized in the thylakoid membranes [43]. Recently, in C. reinhardtii a nuclear-encoded sulfate permease gene (SulP), similar to the CysT gene but with a sequence encoding a transit peptide has been identified [51].

2.2 Sulfate activation (ATP sulfurylase)

The second step in the inorganic sulfate assimilation pathway is catalyzed by ATP sulfurylase (ATPS, EC; Fig. 1, reaction 2). This enzyme activates Embedded Image via an ATP-dependent reaction that leads to the formation of APS and pyrophosphate (PPi). ATPS catalyzes an energetically unfavorable reaction (Δ′=+45.2 kJEmbedded Image[56], an apparently common feature in irreversible reactions under physiological conditions [57]. These thermodynamic and kinetic characteristics make the ATPS reaction a likely rate-limiting step of the SAP.

ATPS of S. cerevisiae and the filamentous fungus Penicillium chrisogenum is an enzyme composed of six identical subunits [58, 59]. The kinetic properties of the S. cerevisiae enzyme are shown in Table 1. In P. chrisogenum, ATPS shows allosteric inhibition by APS and PAPS, which is not observed in the yeast and plant enzymes [59, 60]. In plants, the existence of homodimeric cytosolic and chloroplastic isoenzymes of ATPS has been described [61]; the cytosolic isoform seems involved in a pathway non-related to sulfate reduction, but in production of sulfonated compounds [36, 62]. The ATPS quaternary structure in protists is unknown, but in E. gracilis aplastidic mutants two isoforms localized in cytosol and mitochondria have been identified [63], suggesting that sulfate activation might be performed in both compartments.

Alignment of yeast and plant ATPS gene sequences shows two conserved motifs probably involved in sulfate binding, and a phosphate-binding loop motif, the ATP binding region [64]. In plants, genes encoding cytosolic and plastidic isoforms have been cloned [61, 6568]. In S. cerevisiae, the MET3 gene encodes an ATPS [32]. In C. reinhardtii the ATS1 gene encodes this enzyme [49b], and in protists no information is available.

2.3 Reduction of sulfate to sulfide (APSK, PAPSR, APSR, SiR)

To accomplish the incorporation of sulfur into biomolecules, specifically amino acids, sulfate in APS is transformed to sulfite and this into sulfide. This process may occur through two different pathways, depending on the organism. One of them involves the phosphorylation of APS by an APS kinase (APSK, EC (Fig. 1, reaction 3) using ATP to produce PAPS and ADP. In the following reaction, PAPS reductase (PAPSR, EC 1.8.99) firstly reacts with reduced thioredoxin and then with PAPS to generate free Embedded Image (Fig. 1, reaction 4). The other pathway involves the direct reduction of APS by APS reductase (APSR, EC 1.8.99.x) which uses GSH as an electron source to produce Embedded Image (Fig. 1, reaction 5) [69].

In yeast and many bacteria, Embedded Image is synthesized via APSK [33] whereas in plants, green algae and phototrophic bacteria, sulfate is transformed into sulfite via APSR (for a detailed description of the supporting evidence, see [36, 37]).

APSR activity in Arabidopsis has not been detected in cytosol [62], and the three genes encoding this enzyme contain transit peptides, indicating that this activity is exclusively localized in chloroplasts [70]. Several plant genes encoding cytosolic and chloroplastic APSKs have been cloned [71]. However, it has become clear that PAPS synthesized by APSK is involved in sulfation of several metabolites such as sulfated flavonols, glucosinolates, steroids and phytosulfokines, but not in sulfate reduction and cysteine biosynthetic pathways [36 and references therein].

In S. cerevisiae, MET14 gene encodes APSK, which has a molecular mass of 23 000 [72], and PAPSR is encoded by MET16 [73]. A recent phylogenetic study [74] concludes that the presence of an extra iron-sulfur cluster in APSR determines the enzyme specificity and thus separates the APS- and PAPS-dependent sulfate reduction assimilatory pathways. Since PAPS is a highly toxic compound, the cells must strictly regulate its level. In S. cerevisiae, a 3′(2′),5′-bisphosphonucleoside-3′(2′)-phosphohydrolase encoded by the MET22/HAL2 gene, has been proposed as the enzyme that transforms PAPS into APS to control its intracellular concentration [32]. In P. chrysogenum and Aspergillus nidulans, overaccumulation of PAPS seems to be prevented through PAPS-mediated inhibition of ATPS [75].

The previous idea that plants may reduce sulfate through the PAPS pathway has been re-evaluated with experiments with the knockout of the APSR gene in the moss Physcomitrella patens. The growth and content of soluble thiol-compounds of this knockout are not affected in a medium with sulfate as the sole sulfur source [76]. However, under Cd2+ exposure, growth and content of thiol-compounds of the knockout moss are lower than those of wild type cells. Surprisingly, no PAPSR activity is detected in the APSR knockout moss [76]. These observations suggest that there must be a third pathway for sulfate reduction in P. patens which, however, is unable to sustain an adequate cysteine supply under Cd2+ stress.

A cDNA from the green algae Enteromorpha intestinalis, encoding a plastid APSR, has been cloned and the antibodies produced against the recombinant protein cross-reacted with a 45 kDa polypeptide in several chlorophytes but not in chromophytes [77], suggesting that the APSR is not structurally related between these groups. In protists such as E. gracilis, it is not clear which sulfate reduction pathway is working. APSK activity has been detected in this protist [78] but whether sulfite is produced from APS or PAPS reduction is still unknown.

Once sulfate has been reduced to sulfite, the subsequent step is identical in bacteria, fungi and plants. Sulfite is reduced to sulfide at the expense of oxidizing three molecules of NADPH, by sulfite reductase (SiR, EC; Fig. 1, reaction 6). SiR contains a special acidic heme group called siroheme (redox potential of around −340 mV) and a [4Fe–4S] cluster [79], and catalyzes the reduction of sulfite using electrons donated by ferredoxin. S. cerevisiae SiR (see Table 1 for kinetic parameters) is a hetero-tetramer with a MW of 604 kDa and two types of subunits of 116 and 167 kDa [80]. Although yeast SiR is soluble and therefore considered as a cytosolic enzyme, its intracellular location is not well established. Further work by using cell fractionation or immunocytochemistry may show the enzyme localization. Plant SiR has a molecular mass of approximately 65 kDa, is encoded by only one gene, and is exclusively a chloroplastic enzyme [81]. Genes encoding SiR have been isolated from Z. mays, A. thaliana and N. tabaccum [8183].

2.4 Cysteine biosynthesis (SAT, HAT, OAS/OAH TL, β-CTS and γ-CTL)

Depending on the organism, there are two different ways by which sulfide is incorporated into a carbon backbone to produce cysteine. (1) Sulfide is condensed with O-acetylserine (OAS) by OAS thiol lyase (OAS TL), also called OAS sulfhydrylase, to form cysteine directly [33, 76] (Fig. 1, reaction 10). In this pathway OAS is synthesized by serine acetyl transferase (SAT). (2) OAS TL also catalyzes the condensation of sulfide with O-acetylhomoserine (OAH) to form homocysteine (hCys) (Fig. 1, reaction 7). OAH is synthesized by homoserine O-acetyltransferase (HAT). Then, hCys is transformed into Cys by trans-sulfuration (Fig. 1 reactions 8 and 9), i.e. hCys associates with Ser to form cystathionine (CT) by action of cystathionine β-synthase (β-CTS). CT in turn is dissociated into Cys, α-ketobutyrate and ammonia by cystathionine γ-lyase (γ-CTL) [33, 34, 84, 85]. Cys may also be transformed into hCys by reverse trans-sulfuration catalyzed by cystathionine γ-synthase and cystathionine β-lyase [32, 34]. Kinetic parameters of some of these enzymes are shown in Tables 1 and 2.

View this table:

Kinetic parameters of the enzymes involved in GSH and PCs biosynthesis, and in Cd2+ compartmentation in the vacuole

EnzymeAffinity for different ligands (Km)V m or kcatOrganismReference
γ-ECS, γ-glutamylcysteine synthetase1.4 mM (Glu)13 nmol min−1 (mg protein)−1Candida boidinii[200]
0.4 mM (Cys)
3.1 mM (Ki, GSH)
GS, glutathione synthetase0.27 mM (γ-EC)170 nmol min−1 (mg protein)−1C. boidinii[200]
0.6 mM Gly
PCS, phytochelatin synthasea,b6–13 mM (GSH)ak cat= 0.2 s−1aSilene cucubalusa [113]
b9.2 μM (CdGS2)c0.1 nmol min−1 (mg protein)−1 (rate determined in cell extracts)bA. thalianab [111]
b0.54 μM (Cd2+)
cS. pombec [163]
HMT1, heavy metal transporter 1<30 μM (PC3)?1 nmol min−1 (mg protein)−1S. pombe[22]
YCF1, yeast Cd factor 139 μM (Cd–GS2)15.7 nmol min−1 (mg protein)−1S. cerevisiae[158]
Vacuolar Cd2+ antiporter5.5 μM12.5 nmol min−1 (mg protein)−1S. pombe[159]
  • The enzymes with the most complete kinetic analysis are shown. Superscripts indicate the reference from which the value was obtained.

The SAT pathway, which is used by enteric bacteria, such as Escherichia coli and Salmonella typhimurium [86], and by plants, has been extensively reviewed [84, 87, 88]. Fungi use different cysteine biosynthetic pathways depending on the species. S. cerevisiae uses the CT pathway [32], whereas S. pombe lacks the enzymes for trans-sulfuration but has the enzymes for the SAT pathway [34].

Other fungi such as A. nidulans, Neurospora crassa, Yarrowia lipolytica and Cephalosporium acremonium synthesize cysteine through both the SAT and trans-sulfuration pathways [33]. Similarly, the protist parasite Trypanosoma cruzi possesses the enzymes for both sulfur assimilation and trans-sulfuration and the genes encoding β-CS and SAT have been cloned [89]. A pathway with parallel reactions that generate a critical metabolite (Cys) ensures its constant supply and implies that the enzymes involved are not rate-limiting steps [57].

In plants, OAS TL and SAT are organized as a bienzyme complex called cysteine synthase. These enzyme activities have been found in cytosol, chloroplasts and mitochondria [90, 91], indicating that cysteine may be synthesized in all these compartments (Fig. 1; [84]). The fact that sulfate reduction is only performed in chloroplasts implies that sulfide needs to be translocated from the chloroplast to the other cellular compartments. GSH may mediate such a sulfide exchange since there is a close relationship between the cytosolic and chloroplastic GSH pools [92, 93], and probably also the mitochondrial pool.

2.5 Glutathione biosynthesis (γ-ECS, GS)

In contrast to sulfate reduction, GSH biosynthesis is similar in plants, yeast and protists. GSH is synthesized from cysteine in two consecutive ATP-dependent reactions. In the first step γ-glutamylcysteine (γ-EC) is formed from l-glutamate and l-Cys by γ-glutamylcysteine synthetase (γ-ECS; EC The second step is catalyzed by glutathione synthetase (GS; EC which adds glycine to the C-terminal of γ-EC forming GSH [26] (Fig. 2, reactions 1 and 2; Table 2). Serine and glycine, required for the synthesis of GSH, derive from 3-phosphoglycerate [94].

Although all γ-ECSs catalyze the same reaction and have similar affinities for their substrates, they are not structurally related among kingdoms [95]. By nucleotide sequence comparison, four types of γ-ECSs have been identified. The first class, the animal γ-ECS, is formed by two different subunits, one catalytic and the other regulatory [96]. The remaining classes are monomeric enzymes. The second class, the bacterial enzyme, shows no identity (8%) with the animal catalytic subunit [95]. The third class comprises the yeast (S. cerevisiae and S. pombe) and the protist parasite T. brucei sequences, with an identity of 40–45% with the animal γ-ECS [97]. The fourth class, represented by the plant enzyme, has only 15–19% identity with the animal catalytic subunit. A conserved motif, a putative GSH binding site, is present in all the species analyzed but the structural differences suggest that they have evolved independently [95].

A functional similarity between enzymes from different kingdoms is that GSH may be a strong competitive inhibitor (Ki, 0.1–20 mM) promoting feedback regulation of the pathway [26], which implies that γ-ECS may be rate-limiting of GSH and PCs biosynthesis. Also, l-buthionine-(SR)-sulfoximine (BSO) is a specific and potent inhibitor [98], although the bacterial (E. coli) enzyme is fully inactivated only after longer periods of incubation than those used for the rat kidney enzyme [99, 100]. BSO phosphorylation generates an analog of γ-glutamylphosphate (an intermediate in the catalytic cycle) which binds to the active site producing an irreversible inhibition [98].

GS has been isolated and characterized from different organisms, with their Embedded Image for γ-EC varying between 0.02 and 0.63 mM and the Embedded Image for Gly between 0.3 and 1 mM [26, 101] (Table 2). This enzyme is also present in all organisms but similarly to γ-ECS, it differs structurally among kingdoms [102]. Plant GS genes show high identity with the yeast and human sequences but no homology with bacterial genes, suggesting a divergent origin [103]. GSs of plants and mammals are homodimers with a MW of 56–77 kDa per subunit, whereas the S. pombe enzyme is a hetero tetramer formed by two subunits of 26 and 33 kDa [104]. GS of the protist parasite Plasmodium falciparum is a homodimer, formed by subunits of 77 kDa; its deduced amino acid sequence differs from other GSs particularly at the level of the residues involved in γ-EC binding [103].

2.6 Phytochelatin biosynthesis

Phytochelatins (PCs) are peptides with general formula (γ-Glu-Cys)2–11-Gly synthesized from GSH by phytochelatin synthase (PCS; Fig. 2, reaction 4). PCs have been detected in some yeast [1517, 105], higher plants [20, 106, 107], algae [18], and protists [19, 108], but not in bryophytes [109, 110]. Heavy metals such as Zn2+, Hg2+, Cd2+, Fe3+, Al3+, Cu2+ and Pb2+ act as enzyme activators, Cd2+ being the most potent [12, 14, 111].

PCS is a dipeptidyl (instead of a tripeptidyl) transferase that catalyzes PC chain extension in the C- to N-terminal direction, yielding extended (by one γ-EC unit) PC and Gly, GSH or truncated (by one γ-EC unit) PC [111, 112]. The PCS reaction involves the Cd2+-independent enzyme acylation in one site by GSH and acylation in a second site by a thiol-Cd2+ blocked GSH; these two initial steps generate at least two free Gly molecules. In a third step, the acylated enzyme transfers one γ-EC unit to an upcoming GSH or PC; the other bound γ-EC unit is also released to the medium [111, 112] (Fig. 2, reaction 4). PCS is active as a homo-dimer of 41–50 kDa subunits [111, 114, 115]. Variants of PCs with β-Ala, Ser, Glu or Gln instead of Gly as the ending residue have also been identified in some plants whereas PCs without the C-terminal Gly (desGly PCs) have been found in plants and some yeast [12, 106, 107, 115]. In Glycine max, synthesis of homo-PCs, (γ-Glu-Cys)n-β-Ala, is catalyzed by a specific homophytochelatin synthase (hPCS) [106]. This enzyme is able to use GSH or homoglutathione (hGSH; γ-Glu-Cys-β-Ala) as substrate; however, synthesis of PCs from GSH as the sole substrate is 5-fold more efficient than hPCs synthesis with a GSH/hGSH mixture [106].

PCS genes have been cloned from S. pombe, A. thaliana, Triticum aestivum, Brassica juncea, Thlaspi caerulescens, Glycine max, Athyrium yokoscene and even from the nematode Caenorhabditis elegans [106, 114, 116119]. PCS deduced sequences from fission yeast and plants have similar N-terminal sequences but a less-conserved C-terminal region [14a, 120, 121]. These characteristics, together with the observations that a mutation in A. thaliana (cad1-5) causes premature termination of translation [122] and limited proteolysis of PCS [121] do not prompt total loss of function, have suggested that the N-terminal domain contains the catalytic activity and is presumably highly structured. However, the C-terminal domain also has a role in activity since another A. thaliana mutant truncated in this region presented a Cd2+-sensitive phenotype [121, 122], and decreased thermal stability and responsiveness to heavy metals [121]. It seems that the less-conserved Cys residues, often presented in pairs, together with some Glu residues present in the C-terminal domain have a role as a binding sensor for heavy metals [14a, 120, 123].

PCS might be a regulatory enzyme in PCs synthesis since it is the slowest enzyme in the pathway (Table 2) and in S. pombe and many adult plants Cd2+ exposure does not increase the PCS transcript levels [111, 114, 117] (see also Sections 4 and 5).

The existence of PCs in S. cerevisiae and Neurospora crassa has been demonstrated by mass spectrometry [105]. Since only the shortest phytochelatin (PC-2) is detected, and no PCS gene has been found in these fungi, it has been suggested that PCs may be synthesized by a side reaction of GS [124, 125] and by carboxypeptidase Y [126]. Detection of PCs when only high pressure liquid chromatography with dithionitrobenzene derivatization is used should be viewed with caution since other compounds such as coumarins may yield a false positive signal [109]. Thus, identification of PCs should be established by applying additional methods such as amino acid sequencing and composition [1618, 20] or mass spectrometry [105, 107].

3 Regulation of SAP, Cys and GSH synthesis

3.1 Metabolic regulation

Unicellular and multicellular organisms must be able to maintain a relatively constant intracellular environment. When mild or severe perturbations in the external medium occur, the organisms adjust their internal functions. There is a range of external changes within which the organisms survive, but a large range in which cellular functions are irreversibly impaired. Thus, there must exist metabolic and genetic control mechanisms that regulate the effect of the external changes over the intracellular environment, allowing the organisms to adapt. The cellular control machinery pursues the modulation (activation or inhibition) of critical enzyme activities through (a) short-term (biochemical) mechanisms consisting of non-covalent interactions with some metabolites and covalent enzyme modification and (b) long-term (genetic) mechanisms consisting in the change of the rates of synthesis and degradation of enzymes. A systematic control analysis of the SAP, Cys, GSH and PCs biosynthetic pathways has yet not been carried out. However, several observations concerning the probably predominant control mechanisms have been reported, which are described below.

In S. cerevisiae, sulfate transport is inhibited by internal sulfate and by metabolites derived from sulfate reduction such as APS and cysteine [40]. In addition, OAS increases the activity of all enzymes involved in the SAP [127], whereas cysteine decreases some of them [34, 127]. SAT is strongly inhibited by cysteine [127]. In consequence, if cysteine levels diminish, then SAT is activated, increasing the OAS level and activating sulfate assimilation. When cysteine levels are restored, the SAP enzymes and SAT are again inhibited [128].

In A. thaliana, the cytosolic SAT form is inhibited by low concentrations of cysteine (2–10 μM), but the mitochondrial and chloroplastic isoforms are insensitive, indicating differences in the regulatory mechanisms of cysteine synthesis among organelles [91]. In watermelon, a single change in the C-terminal region (G277C) of a cytosolic SAT isoform, makes the enzyme cysteine insensitive [129].

For GSH synthesis, it has been assumed that feedback inhibition of γ-ECS by GSH is the prime regulation mechanism [26, 101, 130]. In turn, different studies have demonstrated that GSH levels change by modifying γ-ECS activity [92, 131, 132]. Thus, GSH depletion may activate γ-ECS and, hence induce an increase in flux, restoring GSH levels [26, 101, 133]. It should be noted that GSH must be consumed and not only oxidized to overcome the γ-ECS inhibition and thus activate the pathway. For example, when ROS are elevated the GSH/GSSG ratio decreases, but GSH reductase (GR) is usually able to reestablish this ratio to control values, keeping the pathway unaffected (Fig. 2, reaction 3). Moreover, changes in the GSH/GSSG ratio do not affect the transcription levels of γ-ECS and GS [28]. Glutathione S-transferases and PCS (see below) catalyze reactions that consume GSH and in consequence their activation (by increasing availability of their substrates or by overexpression) can overcome the GSH feedback inhibition of γ-ECS resulting in an enhanced pathway flux. Therefore, GSH-consuming enzymes should be also considered when GSH synthesis is analyzed. Cysteine and glycine availability is another mechanism that may contribute to modulate GSH synthesis [29, 92, 134].

3.2 Genetic regulation

Most of the work addressing the transcriptional regulation of genes encoding enzymes of the SAP, and cysteine, GSH, and PC synthesis has focused to the understanding of gene activity under sulfur starvation. However, the response of these genes to Cd2+ exposure has not been fully addressed. In some organisms and depending on the Cd2+ concentration tested, the cysteine and GSH levels may diminish after the first minutes of Cd2+ exposure but hour or days later these levels are restored or, in some cases, enhanced over basal values, suggesting transcriptional activation of SAP and GSH synthesis by Cd2+ [19, 28, 29, 31, 110, 135]. Whether this response is directly related to Cd2+ or to GSH or cysteine depletion induced by Cd2+ stress has not been analyzed. In general, all the genes encoding enzymes of these pathways are transcriptionally up-regulated by sulfur starvation and, in the cases where Cd2+ response has been analyzed, most of them also respond by increasing transcriptional activity.

In S. cerevisiae, SUL1 and SUL2 genes increase their mRNA levels by 9–14 and 5 times, respectively, after Cd2+ exposure [136, 137]. Genes encoding HASTs in plants increase their mRNA levels in response to low sulfate availability [35, 46, 48], and to the internal increase in OAS [138, 139]. Their transcript levels diminish with addition of sulfate, cysteine or GSH to the culture media [47, 48, 139]. In Cd2+-exposed B. juncea roots, but not in leaves, a decrease in a putative low-affinity transporter transcript is observed, suggesting a tissue-specific regulation [140]. In Z. mays, Cd2+ exposure induces an increase in the expression of a HAST gene in roots, which correlates with an enhanced uptake of sulfate [141]. More studies on gene expression and kinetics are required to establish whether sulfate transporters are Cd2+ responsive, particularly in Cd2+ resistant organisms.

Expression of ATPS genes is stimulated by sulfate starvation [31]. Cd2+ exposure induces an increase in the A. thaliana APS3 (ATPS) transcript level (13-fold) [31], in roots and leaves of B. juncea [140], and in the MET3 gene (ATPS) of S. cerevisiae [137, 140]. Cd2+ stress also promotes a 6-fold increase in the ATPS protein level in S. cerevisiae [142], although activity was not determined.

Analysis of S. cerevisiae gene expression in response to 0.03 mM CdCl2, using microarray analysis, determined that MET14 (APSK) and MET16 (PAPSR) increased their expression by 21- and 6-fold, respectively, over the control [136]. The proteome analysis of S. cerevisiae shows that the PAPSR protein increases 5-fold in response to 1 mM Cd2+ exposure [142]. In A. thaliana, APSR transcription increases when the plant grows under sulfur starvation [70, 77]. Genes encoding APSR in B. juncea and A. thaliana (PRH19 and PRH43) are also transcriptionally up-regulated by Cd2+ stress [31, 140].

In the proteome analysis of the S. cerevisiae Cd2+-response, the SiR β subunit increases 2.5 times [142], while in the transcriptome analysis, MET10 (encoding the β subunit) increases its amount of mRNA by 5.5 times [136]. In the presence of 0.2 mM CdCl2 the sir transcript levels increase up to 2-fold [31].

In A. thaliana the OAS TL cytosolic gene Atcys-3A is transcriptionally up-regulated by Cd2+ exposure bringing about an increase in GSH synthesis [29]. In S. cerevisiae, the expression of the CT-γ-lyase gene is increased by 13 times after exposure to 1 mM CdCl2 for 1 h [137]. In protists, there is no information about the SAP genes response to Cd2+ exposure.

The S. cerevisiae GSH1 (γ-ECS) gene is transcriptionally up-regulated by Cd2+ [143] whereas the protein shows a stimulation index of 10 in response to Cd2+ stress [142]. Using the S. cerevisiaeγ-ECS promoter sequence, a 20-fold increase in the associated reporter gene activity was found after its exposure to 0.1 mM CdCl2 [144]. In contrast, Cd2+ does not affect the γ-ECS gene in S. pombe [145]. In plants, Cd2+ induces an increase in the γ-ECS mRNA [28, 29, 146]. Post-translational regulation, such as phosphorylation/dephosphorylation, has been invoked to explain the rapid increase in γ-ECS activity after Cd2+ exposure observed in A. thaliana, without an increase in γ-ECS transcript levels [147]. However, to date no experimental evidence of covalent modulation of γ-ECS has been described.

After Cd2+ exposure, the levels of GS transcripts increase around 2-fold in A. thaliana [28, 31]. These increments are not as strong as those observed for other genes encoding enzymes of the SAP [31]. In S. pombe, Cd2+ does not alter the transcription of the GS gene [145]. In protists, there are no reports about transcriptional regulation of γ-ECS or GS induced by Cd2+ exposure.

PCS is transcriptionally up-regulated (2-fold) after Cd2+ exposure in 5 day-old seedlings of A. thaliana and in T. aestivum roots [148, 116]. In B. juncea leaves, but not in roots, a 4-fold increase in the PCS content was found after Cd2+ exposure, without a concomitant increase in mRNA [117], whereas in S. pombe and mature A. thaliana plants Cd2+ exposure does not affect the transcript level of PCS [111, 114].

Genes encoding SAP enzymes are coordinately regulated in S. cerevisiae. Several regulatory molecules have been implicated in the modulation of the gene expression. AdoMet (S-adenosyl-methionine) was first proposed as the main co-repressor [32]. However, recent data sustain the idea that cysteine is the pathway specific co-repressor for the genes encoding SAP enzymes while OAS is the main co-activator [33, 127, 149]. AdoMet seems to act as a co-factor in the cysteine-mediated repression [149]. Likewise, GSH has a moderate repressive role on specific genes such as MET25 encoding OAS TL, which synthesizes homocysteine, the precursor of cysteine in S. cerevisiae [33]. Based on the finding that cysteine biosynthesis in this yeast is performed by the CT pathway instead of the OAS pathway, the role of OAS in this organism seems to be only as a regulatory element and not as a metabolic intermediate [33].

In plants, an integrative metabolic regulation view emerges from the observation that OAS induces the expression of HAST genes in barley and A. thaliana. OAS also differentially mediates the expression of organ specific isoforms of ATPS, APSR, OAS TL, and γ-ECS in A. thaliana [30, 138, 139]. In turn, cysteine and GSH are known to be negative regulators of gene expression of sulfate transporters, ATPS and APSR [150152]. Thus, OAS, cysteine and GSH levels establish a connection between enzyme activities, metabolic flux and gene expression.

The transcriptional factors involved in the regulation of the genes that respond to Cd2+ stress have not been identified in plants and protists. In S. cerevisiae, the induction of the genes encoding the enzymes of the SAP and GSH biosynthesis by Cd2+ depends upon the transcription factor, Met4p, which is recruited by Met31p, Met32p and Cbf1p, to form a transcriptional complex involved in the activation of most of the methionine biosynthetic genes [153155]. The Yap1 factor, which mediates the oxidative stress response, is also involved in the Cd2+ response [142, 154]. Mutants that do not express this factor (Yap1Δ) are hypersensitive, whereas strains overexpressing the factor are hyper-resistant to Cd2+ [156, 157]. Given the importance that the detoxification systems play in the Cd2+ response, it has been suggested that the YCF1 and gsh1 genes, which encode proteins responsible for sequestration of Cd2+ in the vacuole (see below), are the primary targets by which Yap1 exerts a control of Cd2+ resistance [142]. The Skn7 transcription factor, which cooperates with Yap1 to activate the hydrogen peroxide response, appears to negatively act in Cd2+ response, as a repressor of several genes [156].

Functional proteomic studies in S. cerevisiae show that many proteins change in response to Cd2+, particularly enzymes of the SAP and heat shock proteins. Some others are oxidative stress enzymes (catalase T, thioredoxin, thioperoxidase, Mn-superoxide dismutase, Cu/Zn superoxide dismutase, alkylhydroperoxide reductase), proteases and enzymes from carbohydrate metabolism not related with stress responses [137, 142]. These latter enzymes are isoforms with lower sulfur content that are synthesized to replace existing enzymes. In control conditions the amount of sulfate incorporated into these proteins is 79% but diminishes to 19% under Cd2+ stress. Under this condition, 70% of sulfate is incorporated into metabolites of the GSH biosynthesis (CT, γ-EC and GSH). Thus, the sulfur amino acids may be directed to the massive production of GSH. The fact that the “new enzymes” have less sulfur amino acids, also makes them less susceptible to the deleterious Cd2+ effect since Cd2+ will bind them with a much lower affinity. Based on the linear correlation found between the induction factor of transcript levels and the induction factor of protein content, it was concluded [137] that the yeast Cd2+ response is essentially regulated at the transcriptional level. However, this might be an overinterpretation since no determination of enzyme activity, flux, and time-dependent metabolite variations of the SAP, Cys and GSH pathways were made.

4 Mechanisms of GSH- and PC-mediated Cd2+ resistance

4.1 Yeast and plants

Compartmentation in the vacuole appears to be the most important mechanism for Cd2+ resistance in S. cerevisiae, S. pombe, Candida glabrata, and plants [12, 17, 22, 23, 158, 159]. Cadmium can be transported into the vacuole as a free ion or associated with thiol-compounds (GSH or PCs) [22, 23, 158, 159]. In S. pombe, C. glabrata, some algae and plants, PC–Cd complexes, free Cd2+ and sulfide form high molecular weight (HMW) complexes inside the vacuole which are the ultimate and more stable storage of Cd2+ inside the cell (Fig. 3(a)) [17, 160163]. It should be noted that the available Km values of the transport systems for free Cd2+ (see Table 2) are relatively high (>30 μM), whereas the Kd value of PCS for free Cd2+ is low (0.54 μM). Although no data on free Cd2+ concentrations have been described in organisms exposed to toxic CdCl2 concentrations, the Km and Kd values for Cd2+ may be used as a reference for the expected range of free Cd2+ concentrations that can be reached in the cytosol to turn on the cellular Cd2+ response.


Mechanism of Cd2+ resistance in plants, yeast and protists. (a) Cd–PC complexes are compartmentalised into vacuoles or tonoplasts. In yeast, HMT1 is the protein responsible for transport of the Cd–PC complex [22]. In plants, a similar activity has been described but no homologous gene has yet been found [23, 167]. (b) Proposed mechanism for Cd2+ resistance in E. gracilis, in which most of the Cd2+ and PCs are located inside the chloroplast [19]. Numbered reactions are described in the text.

In S. pombe, PCs are transported into the vacuole by an ABC-type transporter, encoded by the hmt1 gene [22, 164] (heavy metal tolerance 1; Fig. 3(a), reaction 1). The HMT1 protein is related to multi-drug resistance proteins (MRPs), which catalyze the ATP-driven transport of GSH S-conjugates for xenobiotic detoxification. However, unlike MRPs, HMT1 has only one transmembrane and one nucleotide-binding domain [164]. HMT1 activity is sensitive to vanadate, but not to inhibitors affecting the vacuolar H+-ATPase or to ionophores that abolishes the pH gradient across the vacuolar membrane [22], indicating a strict dependence on ATP hydrolysis, but not on a H+ gradient, for driving uptake of Cd2+-PCs complexes.

HMT1 is specific for PC transport and hmt1 cells are Cd2+ sensitive, which indicates that the resistance mechanism in S. pombe depends on the proper storage of PCs in vacuoles. HMT1 gene expression is not Cd2+ inducible, although its overexpression enhances Cd2+ accumulation and resistance [22].

In S. cerevisiae, Cd2+ is also stored into the vacuole as a complex, but in contrast to S. pombe and plants, it is transported as Cd-bis(glutathionate) (Cd–GS2) by YCF1 (yeast cadmium factor) [158, 165] (Fig. 3(a), reaction 2). YCF1 shares amino acid sequence similarity with MRPs and HMT1, its reaction is ATP-dependent and uncoupler-insensitive, but in contrast to hmt1, ycf1 expression is up-regulated when cells are exposed to Cd2+ [158]. YCF1 has a KmCd–GS2 of 39 μM and Vm of 15.7 nmol min−1 (mg protein)−1 [134]. YCF1 may transport Cd–GS2 and some GSH-conjugates such as dinitrophenyl-GSH but no PCs [158]. HMT1 may transport PCs and Cd–PC complexes but apparently no other GSH-conjugates [22]. Recently, another MRP named BPT1 (from bile pigment transporter) has also been related to Cd2+ resistance in S. cerevisiae [166], although its activity does not replace YCF1 function. In plants, PCs are also transported to the tonoplast in an ATP-driven process similar to that reported in S. pombe [23]. However, no hmt1 or ycf1 homolog genes have yet been identified [167].

Other vacuolar membrane transporters in S. pombe, S. cerevisiae and in plants can also take up free Cd2+. Uptake of Cd2+ is not affected by vanadate but it is inhibited by Embedded Image and does not occur when a non-hydrolyzable ATP analog (AMP-PNP) is used. This suggests that Cd2+ transport is driven by the Δ pH generated by the vacuolar H+-ATPase [159, 168] (Fig. 3(a), reaction 3). In Avena sativa vacuoles, the nigericin-dependent Cd2+ uptake has a KmCd of 5.5 μM and Vm of 12 nmol min−1 (mg protein)−1, whereas in S. pombe vacuoles, the initial Cd2+ uptake rate with 8 μM CdCl2 is 37 nmol min−1 (mg protein)−1 [22, 159]. It is not known whether in the cytosol of Cd2+-exposed cells, the free Cd2+ concentration may reach values of 5–8 μM or higher. Due to the tight Cd2+ binding to a variety of biomolecules, including thiols, it is likely that the free Cd2+ concentration be lower than 10 μM under exposure to non-toxic CdCl2 concentrations. A non-manageable level of intracellular free Cd2+ is reached when the cellular chelating capacity is surpassed, which may bring about the usually observed deleterious effects of Cd2+ toxicity on cell physiology.

Five protein families have been implicated in the transport of Cd2+ through cell membranes: (1) cation/H+ antiporter family; (2) CPx-type ATPases; (3) Nramp, natural resistance-associated macrophage proteins; (4) CDF, cation diffusion facilitator family, also named cation-efflux family; (5) ZIP, ZRT-IRT-like proteins (for a detailed description of these families, see [169171]).

In S. cerevisiae, Nramp, CDF and ZIP proteins are responsible for Cd2+ trafficking between cytosol and vacuole. SMF1, one of the three Nramp members identified in S. cerevisiae, is able to take up Cd2+, Cu2+ and Mn2+ [172]. SMF1 is located in the vacuole, although when cells are grown in the absence of heavy metals, it may also be located in the plasma membrane [173].

The S. cerevisiae ZRC1 and COT1 proteins, members of the CDF family, have been related to detoxification of heavy metals [168, 174177]. Both proteins are responsible for Zn2+ storage in the vacuole, but at Zn2+-limiting conditions, ZRC1 is more important than COT1 for metal storage [177]. Cells deleted for either or both genes encoding these proteins show a decreased resistance to Co2+, Zn2+ and Cd2+, with the cot1 deletion being more detrimental to Cd2+ exposure than the zrc1 deletion [177, 178]. These observations suggest that both proteins are involved in the uptake of Cd2+ into the vacuole and that they have different affinities for heavy metals or have different expression in response to heavy metal exposure.

The S. pombe genome has revealed three CDF sequences, and the deletion of one of them (ΔSpZRC1) also results in extremely high Zn2+ and Co2+ sensitivity [178]. The only ZIP family member found in S. cerevisiae (ZRT3) seems to play a role different to heavy metal detoxification since it releases Zn2+ from the vacuole; ZRT3 plays a crucial role when cells are grown in Zn2+-limited medium [176]. All the transporters described above are transcriptionally regulated by Zn2+ availability in the growth medium [176, 177]. Overexpression of ZRC1 and COT1 increases Zn2+ resistance, whereas ZRT3 overexpression results in poor growth [174176]. Unfortunately, neither the response of these transporters to Cd2+ exposure nor the effect of their overexpression on Cd2+ resistance has yet been explored.

In plants, proteins of the cation/H+ antiporter, ZIP, Nramp and CDF families transport Cd2+ (see 171 and references therein). To date, only CAX2 (a cation/H+ antiporter) from N. tabacum has proven to be located in vacuole membranes [179]. CAX2 is not up regulated by CdCl2 exposure, but heterologous expression of A. thaliana CAX2 in tobacco plants leads to 15% more Cd2+ accumulation than that attained in control plants and to a 1.6- to 3-times increase in the Cd2+, Mn2+ and Ca2+ uptake in isolated tonoplast vesicles [179].

In summary, Cd2+ can enter vacuoles in two ways: as a free ion, or complexed with GSH or PCs. In S. pombe, C. glabrata and plants, Cd2+–PC complexes incorporate sulfide to form HMW complexes (Fig. 3(a), reaction 4) around a CdS crystallite core [16, 17, 160163]. The vacuolar S2−/Cd2+ ratio varies between 0.16 and 2 [17, 160, 180]. The addition of sulfide confers a higher Cd2+-binding capacity and enhanced stability to the complex, bringing about full inactivation of the toxic Cd2+ ion. Impairment in any of these processes, i.e. sequestration, transport, and HMW complex formation, results in a Cd2+-hypersensitive phenotype [160, 161, 164].

Studies in C. glabrata have shown that the molecular weight and peptide composition of HMW complexes may vary depending on the sulfide content and the growth media composition [17, 181]. Cells grown in synthetic complete medium form HMW complexes lower than 50 kDa whereas additional sulfur supply at the late logarithmic phase of culture (from 0.2 to 2.1 mM of methionine) gives rise to higher than 50 kDa complexes [181]. In addition, C. glabrata cultured in rich medium contains the usual CdS cores coated with GSH and γ-EC, whereas when it is cultured in minimal medium the CdS core is coated by PC2 and desGly PC2 [182]. The factor involved in this difference is not known; however, it is clear that γ-EC/GSH coated complexes are less stable than those formed by PCs [182].

HMW complexes can be formed in vitro by mixing sulfide, Cd and PCs [165]. However in S. pombe, two purine biosynthetic enzymes, adenylosuccinate synthetase and succinoaminoimidazole carboximide synthetase, are required for HMW complex formation [160, 161]. It is suggested that these enzymes use a sulfur analog of aspartate, cysteine sulfinate, to produce intermediates or carriers in sulfate assimilation to form HMW complexes. However, the reaction sequence in vivo has not been fully elucidated [161].

4.2 Euglena and unicellular green algae

Euglena gracilis is a photosynthetic protist with high Cd2+ tolerance and high Cd2+ accumulating capacity, in which the effects of Cd2+ exposure have been more extensively studied. Probably due to the absence of a specialized reservoir organelle such as a plant-like vacuole, in Cd2+ exposed light-grown cells more than 60% of the accumulated Cd2+ resides inside the chloroplast [19], whereas in dark-grown cells most of it is located in mitochondria [108].

Several reports have shown that the SAP, including GSH and PCs synthesis, is related with the Cd2+ resistance mechanism in this protist [18, 19, 108, 183]. When photosynthetic E. gracilis cells are exposed to 0.2 mM CdCl2, their cellular levels of cysteine, GSH and PCs increase, being 12-fold larger than in non-exposed cells. These metabolite variations occur in the cytosol and more remarkably in the chloroplast [19].

The mechanism of how PCs are synthesized and stored in E. gracilis is under investigation in our laboratory. In this protist, PCs are found in the cytosol, chloroplasts and even mitochondria after Cd2+ exposure [19, 108]. These findings may be explained by either of the following mechanisms. (1) PCs are synthesized in the cytosol where they sequester Cd2+; the Cd–PC complexes are subsequently transported into the chloroplast and mitochondria. (2) PCs are synthesized inside the organelles where they bind Cd2+, which is transported as a free ion, and form HMW complexes. (3) Both processes co-exist and PCs are synthesized in the three cellular compartments (Fig. 3 (b)).

The first possibility implies the existence of an ABC-like transporter to mobilize Cd–PC or Cd–GS2 into the mitochondria and chloroplasts of E. gracilis, similar to that present in S. pombe and plant vacuoles; however, this kind of transporter has not yet been described for chloroplasts or mitochondria.

The second possibility implies the presence of a free Cd2+ transporter in the organellar membranes and a PCS activity inside the organelles. Indeed, E. gracilis chloroplasts are able to take up Cd2+ in its free form. This process involves at least two components [19], one saturable (Vm= 11 nmol min−1 mg protein−1 and Km= 15 μM) and a non-saturable one (0.12 nmol min−1 mg protein−1μM−1) (Fig. 3 (b), reaction 5). In addition, net PCs synthesis by Percoll-purified chloroplasts incubated with 0.5 mM CdCl2 (0.15 nmol h−1 mg protein−1) suggests that a PCS isoform is localized inside E. gracilis chloroplasts. Moreover, we have detected HMW complexes in chloroplasts isolated from Cd2+-exposed Euglena (Mendoza-Cózatl D, Rangel-González E, Moreno-Sánchez R, unpublished data). These observations support the existence of the second mechanism in Euglena chloroplasts, but they do not exclude the existence of the first possibility.

It is difficult to understand how a Cd2+ resistance mechanism might be biochemically and structurally supported in either chloroplasts or mitochondria, which perform several essential reactions and provide energy for the cell function. Hence, Cd2+ accumulation inside these cellular compartments might be a consequence of the organelle characteristics in this particular protist rather than a specific mechanism for Cd2+ resistance. Cd2+ enters the chloroplasts, probably using the transport systems for essential ions. Thus, it seems likely that in the chloroplast mechanisms to maintain low levels of internal free ion metals (essential and non-essential) have evolved to prevent their toxic effects.

PCs have been described in several groups of algae including chlorophytes, xanthophytes, diatoms, phaeophytes and rhodophytes [18]; but there is no information about the detailed function of these peptides or whether they are involved in Cd2+ transport to intracellular organelles. In addition to E. gracilis, it has been found that 60% of accumulated Cd2+ resides inside the chloroplast in a cell-wall deficient strain of C. reinhardtii [184]. HMW complexes are also found in this organelle [184], but the origin of plastid PCs and the mechanism by which Cd2+ is transported into the chloroplast of Chlamydomonas is still unknown. Chlorella synthesizes PCs and forms HMW complexes in response to Cd2+ exposure, although the intracellular localization of these compounds is also unknown [11, 18]. However, it has been established for Chlorella that the cell wall represents the first line of defense against Cd2+ exposure. In fact, at 100 μM CdCl2, 50% of the cellular Cd2+ is bound to the cell wall [185].

Increased PCs synthesis has also been associated with higher resistance to oxidative stress. In the green algae Dunaliella tertiolecta, an increase in PCs induced by Zn2+ exposure, results in an enhanced resistance to ROS caused by H2O2 and paraquat [186]. This protection was the result of a stronger reaction of H2O2 with PC3 than with GSH or ascorbate. Pre-treatment of D. tertiolecta with Zn2+ brings about an increased resistance to Cd2+, through a PC-mediated process [186].

Chlorella vulgaris and some plants accumulate proline when exposed to heavy metals [187, 188]. Because proline may directly react with free radicals diminishing the damage by oxidative stress [189], it was proposed that organisms with high levels of proline might show a higher Cd2+ resistance than organisms with low proline levels [188]. This hypothesis has been tested by using transgenic C. reinhardtii cells overexpressing Δ1-pyrroline-5-carboxylate synthetase, the enzyme that catalyzes the first step in proline biosynthesis in plants. The transgenic cells contained 80% more proline than non-transformed cells. Under Cd2+ exposure, transformed cells showed a more vigorous growth, 4.1 times more Cd2+ accumulation, 80% more GSH, apparently more PCs and a decrease in the GSSG levels. GSH contends simultaneously with Cd2+ detoxification and oxidative stress induced by Cd2+. Therefore, it appears that in the presence of high proline, GSH availability for PCs synthesis is elevated, resulting in a more Cd2+ resistant organism [188].

5 Enhancing glutathione and phytochelatin synthesis

It is desirable that organisms designed for bioremediation, or for enhancing the content of essential metals in food crops, must have some, if not all, of the following characteristics. (1) High heavy metal uptake rate; (2) an efficient mechanism for metal sequestration-inactivation; (3) appropriate heavy metal storage; (4) large biomass production; and (5) in the case of plants, an adequate root-to-shoot transport of the metal. The first point, related to the uptake of the heavy metal from the environment, is probably the process for which modification may be more critical due to the dependence of the subsequent processes on this initial event. In the last decade, the cloning of the genes involved in the SAP, GSH-PCs synthesis, and in intracellular heavy metal and Cd-complex transport has prompted several groups to improve the heavy metal resistance-accumulation capacity of the cells by overexpressing some of the enzymes involved in such processes.

Although some groups have obtained promising results regarding enhanced GSH biosynthesis and Cd2+ resistance [92, 190193], others have not succeeded [92, 132, 191194]. This is probably because most of the overexpression experiments have increased the amount of only one enzyme, the putative rate-limiting step, without considering substrate availability [194] or product inhibition for the overexpressed enzyme or the side accumulation of toxic intermediaries [132]. Moreover, there are no experimental data on how GSH and PC synthesizing pathways may respond to an increased demand of cysteine and GSH (for instance under Cd2+ stress). Thus, it is not known up to which limit an enzyme activity may be elevated without (a) compromising the cysteine and GSH pools (which are also used for protein synthesis, ROS and electrophilic compounds processing) or (b) provoking the accumulation of reactive intermediaries such as oxidized γ-EC (ESSE) [132]. Furthermore, as discussed throughout this review, heavy metal resistance and accumulation is not related to only one enzyme activity, but it is the result of a complex regulation at the genetic and enzymatic levels of several simultaneous processes (sequestration, transport and storage).

We are aware that simultaneous overexpression of several enzymes may be technically difficult. Then, an appropriate analysis of control of Cys, GSH and PCs synthesis, in both unstressed conditions and during Cd2+ exposure, may lead to the identification of a minimal set of enzymes that need to be modulated for reaching enhanced fluxes or metabolite concentrations. Such a set of enzymes to be modified probably may vary between organisms. However, analysis of the kinetic properties of the enzymes of the yeast pathway (Tables 1 and 2), but also of the plant pathway (data not shown), suggests that γ-ECS and PCS may be the most relevant steps in the control of flux, with ATPS and the Cd–PC (or Cd–GS2) vacuole transporter playing a secondary but still significant role. Modulation of flux towards PCs might also be exerted downstream in the pathway, i.e., sulfide transport and incorporation into HMW complexes, but this has not yet been explored.

PCS meets the requirements for an ideal target to be genetically manipulated. (1) It is a homodimeric enzyme coded by one gene; (2) its product is directly involved in sequestration and transport of Cd2+; and (3) different PCS sequences from several organisms are now available. However, some confounding results raise doubts about PCS as a genetic target. For instance, in A. thaliana PCS overexpression causes Cd2+ hypersensitivity [194]. High expression of C. elegans PCS in a Cd2+ hypersensitive strain of S. pombe (Sp27) is less effective in rescuing the Cd2+ hypersensitivity than low expression of the enzyme [119]. In the case of A. thaliana, the possibility of a limited GSH supply is apparently excluded since the total glutathione content (GSSG + GSH) is the same in control and transformed plants. However, Cd2+ exposure may alter the GSH/GSSG balance and this ratio was not evaluated. Therefore, GSH availability may be partially responsible for the confounding result.

On the other hand, the most successful strategy to increase the GSH content has been the overexpression of γ-ECS [92, 190192]. However, an overwhelming activity of γ-ECS over GS activity may also cause oxidative stress by accumulation of ESSE [132]. Therefore, an adequate increase in PCs synthesis together with a sufficient supply of GSH without exceeding the PCs transport rate nor the GS activity may bring about improved results than those obtained by unrestricted overexpression of just one enzyme. Future work is required to determine the control of cysteine, GSH and PCs synthesis and to establish which set of enzymes need to be modulated to obtain enhanced fluxes and increased metabolite concentrations with a minimum of side-effects.

6 Concluding remarks

  1. A widespread response to Cd2+ stress in yeast, plants and protists is the enhancement in the rate of sulfur assimilation, cysteine, GSH, and in some organisms, PCs synthesis through enzymatic activation and up-regulation of some genes involved in these biosynthetic pathways.

  2. Compartmentation of Cd–PC complexes is not restricted to vacuoles or tonoplasts. In photosynthetic protists and some unicellular green algae, other organelles such as chloroplast or mitochondria may be involved in the Cd2+ resistance mechanism.

  3. If Cd2+ resistance is associated with its intracellular accumulation, then the organism with such abilities may be used in the bioremediation of heavy metal-polluted soils and water bodies.

  4. A systematic study of the control of cysteine, GSH and PCs synthesis is required to elucidate the set of enzymes to be enhanced for improving the Cd2+ resistance-accumulation ability of any given organism. This set of enzymes may vary between species.


The present work was partially supported by Grant CONACyT-México No. 43811-Q.


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View Abstract