Abstract

The perceived importance of tellurium (Te) in biological systems has lagged behind selenium (Se), its lighter sister in the Group 16 chalcogens, because of tellurium's lower crustal abundance, lower oxyanion solubility and biospheric mobility and the fact that, unlike Se, Te has yet to be found to be an essential trace element. Te applications in electronics, optics, batteries and mining industries have expanded during the last few years, leading to an increase in environmental Te contamination, thus renewing biological interest in Te toxicity. This chalcogen is rarely found in the nontoxic, elemental state (Te0), but its soluble oxyanions, tellurite (TeO32−) and tellurate (TeO42−), are toxic for most forms of life even at very low concentrations. Although a number of Te resistance determinants (TelR) have been identified in plasmids or in the bacterial chromosome of different species of bacteria, the genetic and/or biochemical basis underlying bacterial TeO32− toxicity is still poorly understood. This review traces the history of Te in its biological interactions, its enigmatic toxicity, importance in cellular oxidative stress, and interaction in cysteine metabolism.

Introduction

Historical background

Tellurium (Te) was discovered by Müller in 1782 (Dittmer et al., 2003) in work with Hungarian gold mines. The metalloid was named 16 years later after the Latin word for earth, tellus (Weeks, 1956) (tellos in Greek). The initial discovery was not so much a determination of a new element as an exclusion of other alternatives, the last being proof that the element, which Müller had isolated from Transylvanian gold ore, was not antimony. Somewhat surprisingly, this isolation came 35 years before the lighter, sister metalloid selenium's discovery by Berzelius in 1817. Sulfur had been known since ancient times and oxygen was isolated in 1774 (Weeks, 1956). Oxygen, sulfur, selenium, and tellurium are collectively referred to as chalcogens (Fischer et al., 2001).

As is characteristic of most of the metalloids, tellurium's oxyanions are relatively stable. Environmentally, tellurite (TeO32−) is most abundant, with tellurate's low solubility (TeO42−) limiting its concentration in biospheric waters. Elemental tellurium (Te0) is insoluble and found as black deposits in some bacterial-selective growth media (Chasteen & Bentley, 2003; Amoozegar et al., 2008). Gold ores containing Te are calaverite (AuTe2), sylvanite (AgAuTe4), and nagyagite [AuPb(Sb, Bi)Te2–3S6] (Cairnes et al., 1911). Te is often found associated with copper- and sulfur-bearing ores (Pan & Xie, 2001; Whiteley & Murray, 2005) and so Te is most commonly commercially obtained as a byproduct of copper refining (Dittmer et al., 2003). Bismuth/tellurium minerals such as tellurobismuthite (Bi2Te3) or wehrite (Bi2Te2) (Cook & Ciobanu, 2004; Laitinen & Oilunkaniemi, 2005) may figure into the story of ‘bismuth breath’ described below.

The biological interaction with inorganic Te compounds was initially reported by Gmelin, who described the olfactory detection of ‘odorous compounds’ in the breath of animals exposed to inorganic Te (Gmelin, 1824). This report influenced the experimental design by Hansen in the mid-1800s – investigating the garlic-like odor in the breath of humans and dogs exposed to TeO32−– who experimented upon himself by taking 30–80 mg doses of TeO32− and postulated that the smelly gaseous compound emitted was diethyl telluride (Hansen, 1853).

This 19th-century experimental exuberance was repeated by Reisert in his pursuit of ‘bismuth breath’, a ‘disagreeable garlic-like odor’ emanating from humans who have ingested bismuth (Bi) salts (Reisert, 1884). When ingested, purified bismuth nitrate yielded no odor from dosed subjects. However, ingested TeO2 at doses as small as 6 μg produced detectable garlic-like odors from human breath that lasted 30 h. Reisert's analogous As-ingestion experiments self-administered on the author, although producing ‘griping’ stomach pains after 4 days of daily, 12 mg arsenous oxide, produced no garlic-like breath odors, leading to the report that these odors were not caused by Bi, but by traces of Te in Bi samples from Bi2Te3. Reisert reiterated Hansen's suggestion that the gaseous, biologically produced compound was CH3CH2TeCH2CH3, but cogently noted that dimethyl telluride also smelled like garlic. Hofmeister (1894) later also made this connection and claimed dimethyl telluride detection – from biological sources – based strictly on smell.

Bird & Challenger (1939) identified the volatile, strong-smelling compound as dimethyl telluride, CH3TeCH3. Mead & Giles (1901) had reported CH3TeCH3 in the breath of mammals dosed with Te oxyanions and in the breath of a chemist who, synthesizing large batches of TeO2, had apparently inhaled the dust. Inhalation of a large dosage led to depression and long periods of sleep as well as strong outgassing of dimethyl telluride (from breath, sweat, and feces), smells that last for months and, in one case, a year after exposure (Mead & Giles, 1901). The unenviable subject was Victor Lenher at Columbia University in New York, who was engaged in early studies on Te0 (Lenher, 1899, 1900).

Both Klett (1900) and Scheurlen (1900) reported the production of black or gray insoluble Te0 in microorganisms amended with TeO32−. Later, this would be proven using X-ray diffraction analysis (Tucker et al., 1962). This resulted from the biological reduction of soluble Te(IV) to the insoluble, elemental form and led to an interesting use of Te0 formation as a microbiological test: building on Gosio's (1905) work, Corper (1915) produced a rapid test of viability for the organism that caused human tuberculosis. When a sterile sodium tellurite solution was added to cells thought to be viable human tubercle bacilli and incubated at 37 °C, a visible blackening occurred within 30–120 min if the culture was live. Corper (1915) was interested in avoiding expensive work with dead cultures in his work on the biochemistry and chemotherapy of tuberculosis (Chasteen & Bentley, 2003). Alexander Fleming, the discoverer of penicillin in the 1920s, also used tellurite-amended media to differentially isolate bacteria (Fleming, 1942). Davis (1914) and Cavazutti (1921) had noted much earlier that different bacterial species and strains exhibited widely differing resistance to TeO32− exposure. Davis (1914) also suggested using TeO32− for a differential diagnosis.

Assessment of the toxicity of Te-containing compounds was systematically performed over half a century ago, with the general conclusion that the simplest salts containing TeO32− (e.g. Na2TeO3) were more toxic to most organisms than TeO42− (Fleming et al., 1932, 1942; Franke & Moxon, 1936; Cooper & Few, 1952; Schroeder et al., 1967). Munn & Hopkins (1925), who examined silver ammonium tellurite and potassium iodotellurite as bacterial disinfectants, found them to be comparable to silver nitrate, and Frazer (1930) found intramuscular injections of suspensions of Te0 in glucose to be successful in the treatment of human syphilis; however, one substantial drawback was an intense garlic odor in the patients' breath and urine. De Meio (1946) also noted garlic exhalations from rats dosed with Te0 and, subsequently, reported that ascorbic acid administered to both rats and humans dosed with Te0 reduced the garlic breath (De Meio, 1947). Ascorbic acid was used to treat industrial workers (exposed to Te-containing dust) who exhibited ‘chronic constant garlic breath,’ and it was reported that odoriferous symptoms were completely eliminated or greatly reduced by taking daily vitamin C treatments, but the symptoms returned when the doses of vitamin C were halted. De Meio (1947) proposed that ascorbic acid reduced oxidized Te back to Te0 before it entered the biological pathway for methylation.

In 1945, Frederic Challenger published his seminal review entitled ‘Biological Methylation,’ focusing on the biological interaction of organisms with arsenic, selenium, and tellurium, among others (Challenger et al., 1945; Chasteen & Bentley, 2003; Zannoni et al., 2008). The mechanism he proposed for the sequential reduction and methylation of Se, he also proposed for Te. Fig. 1 details those steps, beginning with H2TeO3 and yielding CH3TeCH3. A modification of this mechanism to explain the production of dimethyl selenenyl sulfide, CH3SeSCH3, has also been proposed (Chasteen et al., 1993). Recent detection of CH3TeSCH3, dimethyl tellurenyl sulfide, in the headspace of a bacterial culture amended with sodium tellurite makes this mechanism plausible for Te too (Swearingen et al., 2004).

Figure 1

Challenger's Te reduction and methylation mechanism.

Figure 1

Challenger's Te reduction and methylation mechanism.

It has been proposed that the reduction and methylation of toxic, metalloidal oxyanions is a detoxification mechanism because the volatile end products are less toxic than the initial oxyanions (McConnell & Portman, 1952; Wilber et al., 1980; Frankenberger & Arshad, 2001; Chasteen & Bentley, 2003). In investigations of the Se mechanism, one of the Se intermediates, dimethyl selenone [(CH3)2SeO2], has been synthesized and its toxicity has been measured (Fig. 1). Using bacterial growth inhibition as a means of probing relativity toxicity, (CH3)2SeO2 was found to be less toxic than either selenite or selenate (Yu et al., 1997).

More recently, as Se was recognized as an essential element (Stadtman et al., 1996; Chasteen & Bentley, 2003), workers began to investigate the role of Te in biological systems. Schroeder et al. (1967) reported that a typical human being possessed >0.5 g of Te, mostly in bone, and this exceeds the levels of all other trace elements in humans, except for iron, zinc, and rubidium. In the development of a so-called Biological System of the Elements, Markert (1992, 1994) proposed that Te, long thought to be toxic, will eventually be found to be an essential element in a manner similar to Se. This was more recently echoed by Chasteen & Bentley (2003).

Until the advent of GC-MS, the identity of the garlic-like odor produced by bacterial cultures amended with Te salts was based primarily on the smell itself or by wet chemical tests designed to trap a bacterial-produced gas, derivatize it, and compare the derivative's melting point with a standard (Bird & Challenger, 1939). Fleming & Alexander (1972) used GC-MS to confirm the production of both CH3SeCH3 and CH3TeCH3 by metalloid-amended Penicillium sp.

The enigma of TeO32− toxicity

Although rarely found in nature, the tellurium oxyanion, TeO32−, is highly toxic for most bacteria at concentrations as low as 1 μg mL−1 (Taylor et al., 1999). This figure is even more significant when compared with other metals and metalloids such as selenium, chromium, iron, mercury, cadmium, and copper, which become toxic at concentrations about 100-fold higher than that of TeO32− (Nies et al., 1999). For example, in Escherichia coli, the toxic effects of TeO32− begin at concentrations several orders of magnitude lower than the standard determined for heavy metals that are of public health and environmental concern such as cobalt, zinc, and chromium (Nies, 1999; Harrison et al., 2004a).

During the last few years, Te applications in electronics, optics, batteries, and mining industries have expanded, which has indirectly led to increased environmental Te contamination, allowing the isolation of a number of naturally occurring tellurite-resistant bacteria from clinical (Bradley et al., 1985; Taylor et al., 1999) and environmental samples (Summers & Jacoby, 1977; Tantaleán et al., 2003).

A number of genetic Te resistance determinants (TelR) have been identified in different species of bacteria that can be found in plasmids or in the bacterial chromosome. In general, these determinants mediate TeO32− resistance by an as yet unknown mechanism.

Analysis of the nucleotide and the deduced amino acid sequences of the TelR determinants has shown a considerable degree of diversity that has hampered the proposal of a universal TeO32− resistance mechanism. A few putative TeO32− resistance mechanisms have been proposed to date, but they are not supported by definitive experimental evidence. The proposed mechanisms include direct extrusion of TeO32−, TeO32− conversion to volatile, alkylated forms, and enzymatic or nonenzymatic reduction of TeO32− (Te4+) to insoluble Te0.

With regard to the mechanism of direct extrusion of TeO32−, it became clear that it does not constitute a true resistance mechanism, as a decreased influx or an increased efflux of TeO32− is not responsible for the K2TeO3 resistance of E. coli cells expressing TelR determinants (Turner et al., 1995a).

In general, most microorganisms share the ability to reduce TeO32− (Te4+) to the less toxic, Te0. This results in the generation of black deposits of metallic Te inside the cell. At this point, it is important to emphasize that there is a difference between microbial TeO32− resistance and TeO32− reduction. Several tellurite-sensitive microorganisms, for example E. coli K12, are also able to reduce TeO32− (Summers & Jacoby, 1977; Avazeri et al., 1997). Work by Van Fleet-Stalder et al. (2000), using X-ray absorption spectroscopy, confirmed Se0 production by Rhodobacter sphaeroides. Later, Harrison et al. (2004b) described the production of Te0 and Se0 in Staphylococcus aureus and Pseudomonas aeruginosa. It has been suggested that a flavine-dependent reductase, located at the plasma membrane, could play an essential role in TeO32− reduction (Moore & Kaplan, 1992). Similarly, Chiong et al. (1988) and Moscoso et al. (1998) documented the ability of some Gram-negative and Gram-positive thermophilic bacteria to reduce this toxic salt. Enzymatic activities present in crude extracts of these microorganisms were found to be NAD(P)H-dependent [Fig. 2(2)]. In addition, in vivo and in vitro TeO32− reduction by E. coli dihydrolipoamide dehydrogenase has been demonstrated recently (Castro et al., 2008).

Figure 2

A model that illustrates our vision of the participation of certain enzymes (or some of their products) of the cysteine biosynthetic pathway in K2TeO3 resistance. Cys, cysteine; CysK, cysteine synthase; [Fe–S], iron-sulfur center; GSH and GSSG, reduced and oxidized glutathione, respectively; IscS, cysteine desulfurase; OAS, O-acetyl-l-serine; RSH and RSSR, reduced and oxidized thiol, respectively; SOD, superoxide dismutase; SUMT, S-adenosyl-l-methionine uroporphirynogen III C-methyltransferase; YqhD, aldehyde reductase; ZWF, glucose-6-phosphate dehydrogenase; ACN, aconitase; FUM, fumarase; LpdA, E3 component of the pyruvate dehydrogenase; YggE, antioxidant protein; UbiE, C-methyl transferase; SoxRS, oxidative stress regulon (superoxide sensitive); OxyR, oxidative stress regulon (hydrogen peroxide sensitive). To exert its toxicity, TeO32− must enter the target cell, most probably through the phosphate entry route (1). Part of the incoming TeO32− is reduced by nitrate reductase (2). In addition, reduced thiols (3), catalase (20), dihydrolipoamide dehydrogenase (4), and other unspecific reductases can reduce TeO32−. TeO32− reduction to Te0 generates superoxide; increasing ROS levels trigger oxidative stress (5). ROS increase generates cellular damage (6). Superoxide anion generated during TeO32− reduction damages [Fe–S] centers in proteins and enzymes (aconitase and fumarase) (7). Released Fe can generate hydroxyl radical (OH*) through Fenton or Habèr–Weiss reactions (8). IscS desulfurase participates in the recovery of [Fe–S] centers (9). TeO32− reduction decreases reduced thiols, which are restored at the cost of NADPH (10). CysK contributes to restore the intracellular RSH pool (11). SUMT participates in the biosynthesis of the siroheme prosthetic group of sulfite reductase (12). The TeO32− reduction product (Te0) could be further eliminated as alkylated volatile forms of Te by UbiE methyltransferase (13). Generated OH* causes macromolecular damage, especially to DNA (14). Superoxide can initiate membrane lipid peroxidation and protein oxidation (15,16). YqhD detoxifies the cell from reactive aldehydes derived from membrane lipid peroxidation (17). YggE would decrease superoxide levels (18). Increased SOD levels allow superoxide dismutation (19). H2O2 generated by superoxide dismutation is decomposed by catalase (20). Tellurite-generated stress induces the expression of the SoxRS regulon (21). zwf gene induction by SoxRS would allow restoring the NADPH pool (22).

Figure 2

A model that illustrates our vision of the participation of certain enzymes (or some of their products) of the cysteine biosynthetic pathway in K2TeO3 resistance. Cys, cysteine; CysK, cysteine synthase; [Fe–S], iron-sulfur center; GSH and GSSG, reduced and oxidized glutathione, respectively; IscS, cysteine desulfurase; OAS, O-acetyl-l-serine; RSH and RSSR, reduced and oxidized thiol, respectively; SOD, superoxide dismutase; SUMT, S-adenosyl-l-methionine uroporphirynogen III C-methyltransferase; YqhD, aldehyde reductase; ZWF, glucose-6-phosphate dehydrogenase; ACN, aconitase; FUM, fumarase; LpdA, E3 component of the pyruvate dehydrogenase; YggE, antioxidant protein; UbiE, C-methyl transferase; SoxRS, oxidative stress regulon (superoxide sensitive); OxyR, oxidative stress regulon (hydrogen peroxide sensitive). To exert its toxicity, TeO32− must enter the target cell, most probably through the phosphate entry route (1). Part of the incoming TeO32− is reduced by nitrate reductase (2). In addition, reduced thiols (3), catalase (20), dihydrolipoamide dehydrogenase (4), and other unspecific reductases can reduce TeO32−. TeO32− reduction to Te0 generates superoxide; increasing ROS levels trigger oxidative stress (5). ROS increase generates cellular damage (6). Superoxide anion generated during TeO32− reduction damages [Fe–S] centers in proteins and enzymes (aconitase and fumarase) (7). Released Fe can generate hydroxyl radical (OH*) through Fenton or Habèr–Weiss reactions (8). IscS desulfurase participates in the recovery of [Fe–S] centers (9). TeO32− reduction decreases reduced thiols, which are restored at the cost of NADPH (10). CysK contributes to restore the intracellular RSH pool (11). SUMT participates in the biosynthesis of the siroheme prosthetic group of sulfite reductase (12). The TeO32− reduction product (Te0) could be further eliminated as alkylated volatile forms of Te by UbiE methyltransferase (13). Generated OH* causes macromolecular damage, especially to DNA (14). Superoxide can initiate membrane lipid peroxidation and protein oxidation (15,16). YqhD detoxifies the cell from reactive aldehydes derived from membrane lipid peroxidation (17). YggE would decrease superoxide levels (18). Increased SOD levels allow superoxide dismutation (19). H2O2 generated by superoxide dismutation is decomposed by catalase (20). Tellurite-generated stress induces the expression of the SoxRS regulon (21). zwf gene induction by SoxRS would allow restoring the NADPH pool (22).

Apparently, differences between tellurite-sensitive and tellurite-resistant organisms can be associated with mechanisms that cause oxyanion extrusion or by biochemical modifications different from reduction. In the latter case, the generation of methylated forms of Te and Se has been detected in the headspace of recombinant E. coli strains carrying genes of the Gram-positive bacilli Geobacillus stearothermophilus. Methyl telluride is volatile and therefore would be easily eliminated from the cell (Araya et al., 2004; Swearingen et al., 2006) [Fig. 2(13)].

In any case, little is known regarding TeO32− resistance mechanisms in microorganisms. To date, five genetic TelR determinants have been characterized in Gram-negative bacteria. Interestingly, four of them were found in plasmids, which are important for the transference of resistance determinants between species. The presence of TelR determinants in a wide range of bacterial species, including those pathogenic for humans, suggests that these determinants provide some selective advantage in their natural environment (Walter & Taylor, 1992; Hill et al., 1993). However, these determinants might not have evolved specifically to provide TeO32− resistance. For example, IncHI2 and IncHII conjugative plasmids carrying the ter operon confer high-level TeO32− resistance as well as resistance to bacteriophages and colicins (Whelan et al., 1995; Taylor et al., 1999; Taylor et al., 2002). In this context, besides the genetic variability observed in E. coli O157 clinical isolates, several putative TeO32− determinants have been found in cross searches by homology in other bacteria such as Yersinia pestis and Deinococcus radiodurans (Taylor et al., 1999; Taylor et al., 2002).

As mentioned above, TeO32− resistance determinants found in extrachromosomal elements include IncHI-2 (Whelan et al., 1995, 1997) and pMER610 (Jobling & Ritchie, 1987, 1988; Hill et al., 1993) plasmids. The unique structure of the Klebsiella pneumoniae TerB protein (151 amino acid residues, KP-TerB) has recently been determined (Chiang et al., 2008). Other examples are the kilA operon (klaA klaB telB) from the RK2/RP4 plasmid, involved in plasmid partition and maintenance (Goncharoff et al., 1991; Walter et al., 1991; Turner et al., 1995b), and the ars operon from the E. coli R773 plasmid (Turner et al., 1992). The kilA operon present in Klebsiella aerogenes plasmid pRK2TeR (accession #M62846), pTB11 plasmid (AJ744860), P. aeruginosa pBS228 plasmid (AM261760) and pWFRT-tel (EU329006) and mini-Tn7-tel (EU626136) cloning vectors share 99–100% sequence identity and encode KlaA, KlaB, and KlaC polypeptides of 257, 378, and 317 amino acid residues, respectively. The genetic and protein contexts are identical and genes are apparently transcribed as an operon. Recently, kilA telAB sequences have been used as selection markers to engineer cloning vehicles for Burkholderia spp. (Barrett et al., 2008). Regarding the arsenical efflux pump, it has been proposed that ArsC is involved in modifying the substrate-binding site of the anion-translocating ATPase, thus conferring moderate levels of resistance to TeO32− (Turner et al., 1992).

On the other hand, Taylor et al. (1994) reported that a fifth TelR determinant, represented by the tehAB operon and located near the terminus of the E. coli chromosome, conferred resistance to potassium tellurite in this bacterium, provided these genes are expressed from a multicopy plasmid (Taylor et al., 1994). Later, Turner et al. (1997) demonstrated that the 36-kDa integral membrane protein TehA confers resistance to antiseptics and disinfectants similar to that conferred by multidrug resistance efflux pumps. More recently, the presence of the resistance determinant tehAB, by an as yet unidentified mechanism, was found to protect the cells from uncoupling by TeO32− (Lohmeier-Vogel et al., 2004).

TehA and TehB orthologs have been found in a number of bacteria including K. pneumoniae (accession #YP_002238239 and YP_002238240, respectively), Salmonella enterica serovar Typhi (CAD01716 and CAD01717), Shigella dysenteriae (YP_403356 and YP_403355), Shigella flexneri (YP_689245 and YP_689244), Haemophilus influenzae (YP_248222 and YP_249313), Pasteurella multocida (NP_ 246526 and NP_245593), S. enterica serovar Typhimurium (NP_460568 and NP_460567), Mannheimia succiniproducens (YP_087218 and YP_088530), and Escherichia albertii (ZP_02902738 and ZP_02902769), among others. Recently, the crystal structures of TehB from Vibrio fischeri (PDB ID: 3DL3) and Corynebacterium glutamicum (PDB ID: 3CGG) have been made available.

It is interesting to note that there is little homology among the nucleotide sequences of these five TelR determinants. Apart from arsRDABC, which encodes an oxyanion efflux system, mechanisms by which all other TelR determinants specify TeO32− resistance are a matter of speculation to date.

Using a different approach, a number of research groups have reported that overexpressing some genes involved in basal metabolism results in an increased tolerance to TeO32−. For example, cloned cysK genes (encoding cysteine synthase) from G. stearothermophilus V (Vásquez et al., 2001), S. aureus (Lithgow et al., 2004), E. coli (Alonso et al., 2000), and R. sphaeroides (O'Gara et al., 1997) mediate TeO32− resistance when expressed in heterologous hosts [Fig. 2(11)].

A possible explanation for these findings may lie in the intrinsic mechanism of TeO32− toxicity. It is known that Te (and Se) oxyanions interact with cellular thiols (RSH), and it has been found that glutathione (GSH) is one of the most important targets of TeO32− in E. coli (Spallholz et al., 1994; Turner et al., 2001). Based on the similarity between Se and Te chemistry, it was postulated that GSH can reduce TeO32− to Te0 (Turner et al., 2001).

On the other hand, it was postulated that superoxide could be generated during TeO32− reduction [Fig. 2(5)] as occurs for selenite oxyanions (SeO32−) (Bébien et al., 2002; Kessi & Hanselmann, 2004). Even though this last possibility has been supported in recent communications (Tantaleán et al., 2003; Borsetti et al., 2005; Rojas & Vásquez, 2005; Calderón et al., 2006), direct experimental evidence was only recently obtained for Pseudomonas pseudoalcaligenes KF707 (Tremaroli et al., 2007) and E. coli (Pérez et al., 2007), where the amount of mRNA transcripts of genes specifically commanded by the transcriptional OxyR and SoxS regulators and the activity of classic antioxidant enzymes such as superoxide dismutase and catalase [Fig. 2(19–20)] were found to be increased as a consequence of TeO32− exposure (Pérez et al., 2007). Thus, an emerging view for the TeO32− toxicity problem is that bacteria seem to cope with the toxicity by a general adaptation mechanism like those used when they face other environmental stressors such as UV radiation or heat shock, among others (Fig. 2). A summary of genes known to be involved in TeO32− resistance or sensitivity is listed in Table 1. Another group of metabolic genes that seems to participate actively in TeO32− metabolism includes acnA, acnB, fumA, and fumC, among others (C. Vásquez, unpublished data).

Table 1

Genes involved in tellurite resistance (R) or sensitivity (S)

Gene symbol Function R/S Organism References 
arsABC Metalloid efflux E. coli Turner et al. (1992) 
aceE, aceF Central metabolism E. coli Castro et al. (2009) 
choQ Amino acid transport Lactococcus lactis Turner et al. (2007) 
CSD (cdsACysteine metabolism E. coli Rojas & Vásquez (2005) 
cobA Siroheme biosynthesis E. coli Araya et al. (2009) 
csdB Cysteine metabolism E. coli Rojas & Vásquez (2005) 
cysM Cysteine metabolism E. coli Lithgow et al. (2004) 
cysK Cysteine metabolism E. coli, Rhodobacter sphaeroides, G. stearothermophilus O'Gara et al. (1997), Alonso et al. (2000), Vásquez et al. (2001), Fuentes et al. (2007) 
gutS Tellurite/selenite-induced transporter ND E. coli Guzzo & Dubow (2000) 
ibpA Heat-shock response E. coli Pérez et al. (2007) 
iscS Cysteine metabolism E. coli Tantaleán et al. (2003), Rojas & Vásquez et al. (2005) 
katA Hydrogen peroxide detoxification Staphylococcus epidermidis Calderón et al. (2006) 
katG Hydrogen peroxide detoxification E. coli Pérez et al. (2007) 
kilA, telA,telB (klaA, klaB, klaCTellurite resistance E. coli Goncharoff et al. (1991), Turner et al. (1995b), Walter et al. (1991) 
lpdA Central metabolism A. caviae Castro et al. (2008) 
mntH Mn+2/Fe+2 transport L. lactis Turner et al. (2007) 
narGHIJ Nitrate reduction E. coli Avazeri et al. (1997) 
phoB, phoRPhosphate metabolism E. coli Tomás & Kay (1986) 
pstA, pstD Phosphate transport L. lactis Turner et al. (2007) 
sodAB Superoxide detoxification E. coli Tantaleán et al. (2003), Pérez et al. (2007) 
soxS Oxidative stress response E. coli Pérez et al. (2007) 
tehA, tehB Tellurite resistance E. coli Taylor et al. (1994), Turner et al. (1995b) 
terBCDE Tellurite resistance E. coli Kormutakova et al. (2000) 
terC Tellurite resistance Proteus mirabilis Toptchieva et al. (2003) 
tmp Purine metabolism P. syringae Cournoyer et al. (1998) 
trgAB Tellurite resistance R. sphaeroides O'Gara et al. (1997) 
trmA Heat-shock response L. lactis Turner et al. (2007) 
ubiE Ubiquinone/menaquinone biosynthesis G. stearothermophilus Araya et al. (2004) 
yqhD Oxidative stress response E. coli Pérez et al. (2008) 
Gene symbol Function R/S Organism References 
arsABC Metalloid efflux E. coli Turner et al. (1992) 
aceE, aceF Central metabolism E. coli Castro et al. (2009) 
choQ Amino acid transport Lactococcus lactis Turner et al. (2007) 
CSD (cdsACysteine metabolism E. coli Rojas & Vásquez (2005) 
cobA Siroheme biosynthesis E. coli Araya et al. (2009) 
csdB Cysteine metabolism E. coli Rojas & Vásquez (2005) 
cysM Cysteine metabolism E. coli Lithgow et al. (2004) 
cysK Cysteine metabolism E. coli, Rhodobacter sphaeroides, G. stearothermophilus O'Gara et al. (1997), Alonso et al. (2000), Vásquez et al. (2001), Fuentes et al. (2007) 
gutS Tellurite/selenite-induced transporter ND E. coli Guzzo & Dubow (2000) 
ibpA Heat-shock response E. coli Pérez et al. (2007) 
iscS Cysteine metabolism E. coli Tantaleán et al. (2003), Rojas & Vásquez et al. (2005) 
katA Hydrogen peroxide detoxification Staphylococcus epidermidis Calderón et al. (2006) 
katG Hydrogen peroxide detoxification E. coli Pérez et al. (2007) 
kilA, telA,telB (klaA, klaB, klaCTellurite resistance E. coli Goncharoff et al. (1991), Turner et al. (1995b), Walter et al. (1991) 
lpdA Central metabolism A. caviae Castro et al. (2008) 
mntH Mn+2/Fe+2 transport L. lactis Turner et al. (2007) 
narGHIJ Nitrate reduction E. coli Avazeri et al. (1997) 
phoB, phoRPhosphate metabolism E. coli Tomás & Kay (1986) 
pstA, pstD Phosphate transport L. lactis Turner et al. (2007) 
sodAB Superoxide detoxification E. coli Tantaleán et al. (2003), Pérez et al. (2007) 
soxS Oxidative stress response E. coli Pérez et al. (2007) 
tehA, tehB Tellurite resistance E. coli Taylor et al. (1994), Turner et al. (1995b) 
terBCDE Tellurite resistance E. coli Kormutakova et al. (2000) 
terC Tellurite resistance Proteus mirabilis Toptchieva et al. (2003) 
tmp Purine metabolism P. syringae Cournoyer et al. (1998) 
trgAB Tellurite resistance R. sphaeroides O'Gara et al. (1997) 
trmA Heat-shock response L. lactis Turner et al. (2007) 
ubiE Ubiquinone/menaquinone biosynthesis G. stearothermophilus Araya et al. (2004) 
yqhD Oxidative stress response E. coli Pérez et al. (2008) 

ND, not defined.

Bacterial mechanisms against oxidative stress and TeO32− tolerance

Bacteria have evolved several mechanisms to protect themselves from environmental stress. The increase in reactive oxygen species (ROS) during oxidative stress leads to thiol oxidation, among other effects. Some of these thiols form part of cellular proteins such as the OxyR transcriptional regulator, which is transitorily activated by disulfide linkage formation under oxidative stress (Zheng et al., 1998). In E. coli, OxyR regulates the expression of several H2O2-inducible genes encoding for enzymes known to participate in the bacterial response to oxidative stress (Zheng et al., 2001). Examples of these enzymes are hydroperoxidase I (encoded by katG), alquil-hydroperoxide reductase (ahpCF), glutathione reductase (gorA), glutaredoxin 1 (grxA), thioredoxin 2 (trxC), iron uptake regulator (fur), an unspecific DNA-binding protein (dps), external membrane protein (agn43) and ferric reductase (fhuF), among others (Storz & Imlay, 1999).

In addition, several E. coli genes are regulated by the soxRS transcriptional regulatory system that responds to superoxide-generating species. These genes include sodA (Mn-dependent superoxide dismutase), nfo (endonuclease IV, involved in DNA repair), zwf (glucose-6-phosphate dehydrogenase), tolC (outer membrane protein), fur, micF (regulatory RNA of ompF expression), acrAB (multidrug efflux pump), fumC (fumarase C), acnA (aconitase A), nfsA (nitroreductase A), fpr (ferredoxin/flavodoxin reductase), fldA, fldB (flavodoxin A and B), and ribA (GTP hydrolase) (Liochev et al., 1999; Storz & Imlay, 1999).

In general, exposure to low stress levels or to some chemicals induces an increase in an organism's resistance to subsequent expositions to the same (adaptative response) or to unrelated (cross-response) agents (Mongkolsuk et al., 1997). In this sense, both types of responses (OxyR and SoxRS) have been shown to play an important role in bacterial oxidative stress and in the stress generated by toxic metals as well. For instance, low concentrations of cadmium induce a synergistic protection against death by hydrogen peroxide in Xanthomonas campestris (Banjerdkij et al., 2005). In turn, sublethal concentrations of selenite activate an adaptive response that increases SeO32− tolerance and induces crossed protection to the ROS elicitor paraquat in E. coli (Bébien et al., 2002). Conversely, treatment with sublethal concentrations of TeO32− did not show cross-protection against the oxidant compounds paraquat, diamide or hydrogen peroxide in P. pseudoalcaligenes KF707 (Tremaroli et al., 2007). These results differ from those observed in E. coli K-12. Pérez et al. (2007) found a cooperative-like toxicity effect between TeO32− and these oxidants as well as an activation of the transcription of oxyR- and soxS-regulated genes, suggesting a synergistic protection against these toxic compounds.

Preliminary results from our laboratory indicate that E. coli cells exposed to paraquat exhibit increased minimal inhibitory concentrations of TeO32−, suggesting an adaptative response of the bacterium. In addition, a clinical isolate of Proteus mirabilis showed increased tolerance to potassium TeO32− when previously grown in media containing sublethal TeO32− concentrations (Toptchieva et al., 2003).

A recent report indicated that an aldehyde reductase, YqhD [Fig. 2(17)], is involved in protecting E. coli against lipid peroxidation caused by the oxidative stress elicitors paraquat, H2O2, chromate or TeO32−, and that this enzyme may be part of a glutathione-independent antioxidant system (Pérez et al., 2008). Thus, at least in P. pseudoalcaligenes KF707 and in E. coli K-12, TeO32− toxic effects seem to be directly related to increased intracellular ROS as well as a decrease in the cellular thiol content (Pérez et al., 2007; Tremaroli et al., 2007).

In this context, two main mechanisms controlling the cytoplasmatic redox balance appear to be directly affected by TeO32−. These are the glutathione–glutaredoxin and the thioredoxin systems (Holmgren, 1989; Carmel-Harel & Storz, 2000). The tripeptide glutathione (l-γ-glutamyl-cysteinyl-glycine) and thioredoxin (12 kDa) act as general reducers in the cell's cytoplasm (Prinz et al., 1997; Smirnova et al., 1999). GSH reduces intracellular disulfide bonds, generally in conjunction with glutaredoxin (Carmel-Harel & Storz, 2000). Being predominantly reduced, cytoplasmic GSH (c. 5 mM) is thought to be the main controller of the redox environment in the cytoplasm of E. coli (Aslund & Beckwith, 1999). In turn, reduced glutathione levels are controlled by glutathione reductase, which reduces oxidized glutathione (GSSG) in an NADPH-dependent reaction. The coenzyme is then recycled by glucose-6-phosphate dehydrogenase in the pentose phosphate pathway [Fig. 2(22)].

In spite of the numerous cell processes that involve glutathione, including its role in defense against oxidative stress (Carmel-Harel & Storz, 2000), it has been observed that this molecule does not appear to be essential for E. coli survival (Fuchs & Warner, 1975). In addition, and although common in most Gram-negative bacteria, GSH has been shown to be present in only a few Gram-positive microorganisms (Fahey et al., 1978).

Three glutaredoxins (1–3 encoded by grxA, grxB, and grxC, respectively) and two thioredoxins (1 and 2, encoded by trxA and trxC, respectively) have been described in E. coli. Most glutaredoxins, all thioredoxins, and thioredoxin reductase (encoded by trxB) contain a conserved motif, CXXC, at the active site. They also share similar structures containing three α-helices and four-fiber β-sheets.

Escherichia coli mutations in GSH and glutaredoxin systems have shown that thioredoxin 1 (TrxA), thioredoxin 2 (TrxC), and thioredoxin reductase (TrxB) are not essential for the bacterium. trxA and trxB mutants, a phenotype that was not observed with trxC cells (Takemoto et al., 1998; Ritz et al., 2000), showed increased sensitivity to H2O2.

Albeit viable, E. coli lacking genes involved in GSH biosynthesis such as gshA, gshB, and gorA (encoding glutathione synthetase isoenzymes and glutathione reductase, respectively) are highly sensitive to the specific thiol oxidizer diamide (Li et al., 2003). Under normal growth conditions, these mutants exhibit a phenotype similar to that of the parental strain. Similar results were observed for grxA and grxC mutants (Aslund et al., 1994), suggesting an independent requirement of these two systems to cope with oxidative stress under these culture conditions. In this context, the GSH-dependent protective system is mainly observed in Gram-negative bacteria, with the exception of Lactococcus lactis and some Bacillus and Clostridium species (Fahey et al., 1978; Zhang et al., 2007).

Double gsh/grx E. coli mutants revealed that the integrity of at least one of these systems is required for aerobic growth of this bacterium (Prinz et al., 1997). These mutants also proved to be unable to reduce disulfide linkage in the cytoplasm. Although these observations emphasize the importance of GSH in maintaining the cytoplasm's reducing atmosphere, the idea is that this kind of function would not be represented exclusively by these reductants in bacteria. This has prompted workers to pursue a continuous search of alternative reducing systems that may involve other thiol-containing molecules.

Sulfur and cysteine metabolism and TeO32− tolerance

Living organisms require sulfur for the synthesis of proteins and essential cofactors. Bacteria and plants acquire sulfur through assimilation from inorganic sources such as sulfate and thiosulfate, or from organic ones such as sulfate esters, sulfamates, and sulfonates (Uria-Nickelsen et al., 1993).

One of the main products of sulfur assimilation is the amino acid cysteine, which performs vital functions in enzyme catalysis and in protein structure, among others. For example, cysteine residues play a crucial role in [Fe–S] cluster-containing proteins as cytochromes and dehydratases of intermediary metabolic pathways such as fumarases and aconitases [Fig. 2(7)]. Important Cys residues have also been described in E. coli Te resistance proteins TehA and TehB. Replacement of these cysteines by alanine causes decreased TeO32− resistance (Dyllick-Brenzinger et al., 2000).

Previous work with G. stearothermophilus and S. aureus has pointed out the relevance of cysteine-metabolism-related genes and K2TeO3 resistance. Three genes coding for proteins acting in sulfur assimilation or using cysteine as a substrate have been identified in G. stearothermophilus: cobA, encoding SUMT methyltransferase [Fig. 2(12)], an enzyme that participates in the biosynthesis of siroheme, a cofactor of sulfite reductase (Araya et al., 2009), cysK, encoding cysteine synthase [Fig. 2(11)], which catalyzes the last step in cysteine biosynthesis (Vásquez et al., 2001; Saavedra et al., 2004), and iscS, specifying the IscS cysteine desulfurase [Fig. 2(9)], which is involved, among other functions, in the recovery and synthesis of [Fe–S] cluster-containing enzymes (Flint et al., 1996; Tantaleán et al., 2003). Recent publications have shown the involvement of cysK and cysM genes in K2TeO3 resistance as well as in response to oxidative stress (Lithgow et al., 2004; Das et al., 2005) and that cysK is differentially expressed in response to H2O2, paraquat, and diamide (Pomposiello et al., 2001; Leichert et al., 2003). Much earlier, cysK mutants of E. coli had also been obtained from selenite-resistant mutants (Fimmel & Loughlin, 1977).

It has long been argued that TeO32− toxicity could be a consequence of its strong oxidant nature (Siliprandi et al., 1971; O'Gara et al., 1997; Garberg et al., 1999; Taylor et al., 1999). Exposure would eventually result in an oxidative stress status in the cell. This last condition is associated with the presence of ROS such as hydrogen peroxide, superoxide, and hydroxyl radicals. In this context, intracellular enzymatic TeO32− reduction by nitrate reductase (Avazeri et al., 1997), dihydrolipoamide dehydrogenase [Castro et al., 2008; Fig. 2(4)], catalase, or by other enzymes [Fig. 2(2)] would result in the generation of superoxide (Calderón et al., 2006; Pérez et al., 2007). TeO32− could also be reduced chemically to lower oxidation states by glutathione or by other reduced thiol-containing molecules [Fig. 2(3)]. A direct consequence of this reduction reaction would be a drastic decrease in the concentration of antioxidant molecules such as glutathione and cysteine, among others. Thus, it would be possible to speculate that turning on the biosynthetic machinery of cellular antioxidants would result in a phenotype of higher TeO32− tolerance.

Recent research described a Gram-negative bacterium, which, in part, responded to TeO32− by conversion to Te0, which was found to be deposited in the cytoplasm (Pages et al., 2008). This microorganism could grow in the presence of 25 mM TeO32− and 50 mM selenite, another toxic oxyanion, which was also reduced and collected in the cell as a precipitated metalloid. A survey of Gram-positive bacteria and yeasts isolated from salt marsh sediments that were TeO32− resistant produced reduced volatile organo-tellurides, mostly dimethyl telluride, when amended with TeO32−. Some strains, when exposed to metalloidal salts, exhibited intracellular elemental Te precipitates (Ollivier et al., 2008).

On the other hand, E. coli cells expressing defined G. stearothermophilus genes (cloned in low copy number plasmids along with their own promoters) exhibit increased tolerance to potassium tellurite and to some oxidative stress elicitors along with some protection against the poisonous effect of diamide, a general thiol oxidizer (Fuentes et al., 2007). Similar results were previously reported when the Bacillus subtilis gene expression profile was analyzed after growing in the presence of hydrogen peroxide (Leichert et al., 2003). More recently, it was shown that TeO32− activates superoxide dismutase in P. pseudoalcaligenes (Tremaroli et al., 2007) and appears to induce the expression of some genes of the OxyR and SoxR regulons of E. coli (Pérez et al., 2007). Moreover, it was shown that TeO32− positively regulates gutS and the genes of the terZABCDE operon in E. coli and P. mirabilis, respectively (Guzzo & Dubow, 2000; Toptchieva et al., 2003). The presence of sequences similar to the OxyR-binding motifs in the operon of P. mirabilis suggests that its induction by TeO32− would be dependent on this transcriptional regulator.

Finally, most genes belonging to the E. coli Cys regulon were shown to be induced in the presence of potassium tellurite, even though cysB and cysE expression was repressed when cells were grown in rich media (Fuentes et al., 2007). In this context, E. coli strains lacking genes involved in cysteine biosynthesis exhibit an increased sensitivity to K2TeO3 (C. Vásquez, unpublished data), suggesting that most components of cysteine metabolism are required to cope with the stress caused by TeO32−. These results show that cysteine-metabolism-related genes are induced upon TeO32− exposure and that cysteine, sulfide, or thiosulfate repression is probably due to a strong intracellular reduction of reduced thiols. Thus, a direct relationship between cysteine biosynthesis and TeO32− tolerance can be established in both Gram-negative (E. coli) and Gram-positive (G. stearothermophilus) bacteria. In support of this, recent evidence by Tremaroli (2009) showed that high TeO32− resistance in P. pseudoalcaligenes KF707 can be correlated with a reconfiguration of the cellular metabolism as well as with an induced oxidative stress response.

Concluding remarks

Although a necessary biochemical role for Te has not been established as it has for Se, it is not unreasonable to assume that ultimately Te will be found to be a necessary trace metalloid for some organisms.

Although the reduction and methylation mechanism has not yet been determined, the production of insoluble Te appears to be a common means of partially detoxifying much more damaging and highly oxidative Te oxyanions.

Why is TeO32− so toxic? What are tellurite's main intracellular targets? What other thiol pool in addition to GSH, if any, is depleted upon TeO32− exposure? Why does the cell assume the apparently unnecessary risk of superoxide generation upon TeO32− reduction? What is the ultimate mechanism underlying TeO32− resistance/toxicity?

In spite of the fact that progress in the field is slow and more effort is needed, the answer to these intriguing questions in the near future will help us to better comprehend the complex phenomena of TeO32− toxicity and bacterial TeO32− resistance.

Acknowledgements

The authors would like to thank Dr Simon Silver (University of Illinois, Chicago) for critically reading the manuscript. This work received financial support from Fondecyt grant # 1060022 and Dicyt-USACH to C.C.V. and from the Robert A. Welch Foundation (X-011) at Sam Houston State University to T.G.C.

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