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The Confluence of Heavy Metal Biooxidation and Heavy Metal Resistance: Implications for Bioleaching by Extreme Thermoacidophiles

Department of Chemical and Biomolecular Engineering, North Carolina State University, EB-1, 911 Partners Way, Raleigh, NC 27695-7905, USA
Author to whom correspondence should be addressed.
Present address: Alexion Pharmaceuticals, 352 Knotter Dr, Cheshire, CT 06410-1138, USA.
Minerals 2015, 5(3), 397-451;
Submission received: 8 May 2015 / Revised: 23 June 2015 / Accepted: 26 June 2015 / Published: 7 July 2015
(This article belongs to the Special Issue Advances in Biohydrometallurgy)


Extreme thermoacidophiles (Topt > 65 °C, pHopt < 3.5) inhabit unique environments fraught with challenges, including extremely high temperatures, low pH, as well as high levels of soluble metal species. In fact, certain members of this group thrive by metabolizing heavy metals, creating a dynamic equilibrium between biooxidation to meet bioenergetic needs and mechanisms for tolerating and resisting the toxic effects of solubilized metals. Extremely thermoacidophilic archaea dominate bioleaching operations at elevated temperatures and have been considered for processing certain mineral types (e.g., chalcopyrite), some of which are recalcitrant to their mesophilic counterparts. A key issue to consider, in addition to temperature and pH, is the extent to which solid phase heavy metals are solubilized and the concomitant impact of these mobilized metals on the microorganism’s growth physiology. Here, extreme thermoacidophiles are examined from the perspectives of biodiversity, heavy metal biooxidation, metal resistance mechanisms, microbe-solid interactions, and application of these archaea in biomining operations.

1. Introduction

The commercial application of microorganisms for the extraction of metals from sulfide ores, concentrates, low-grade ores and tailings, often referred to as bioleaching and biomining, falls within the discipline biohydrometallurgy [1,2]. Bioleaching leverages microbially-based conversion of insoluble metal sulfides (or oxides) to water-soluble metal sulfates. For example, conversion of insoluble chalcopyrite (CuFeS2) to a soluble copper sulfate has become the basis for technologically important processes. Similarly, microorganisms have also been used as a pretreatment step for metals recovery. For example, the removal of sulfur from sulfidic gold ores can enhance downstream recovery and limit depletion of the cyanide extraction agent [3]. The development of bioleaching and biomining technologies has been ongoing for several decades, and more recently is finding increased interest in commercial application. Advances in molecular microbiology and genomic sciences present new opportunities for discovering, characterizing and implementing microbial systems for the recovery of base, precious and strategic metals.
The future of biomining was once declared to be “hot”, owing to the recalcitrance of metal sulfides such as chalcopyrite at moderate temperatures, thereby requiring thermal conditions (65–80 °C) to obtain increased solubilization rates [4,5,6]. At higher temperatures, metal sulfide mobilizing consortia are dominated by archaea, mainly belonging to the genera Acidianus, Metallosphaera, and Sulfolobus [7,8,9,10,11]. The mechanisms by which metal biooxidation, metal resistance, and microbe-solid interactions take place in thermal, acidic environments are not well understood but, if elucidated, could provide valuable information necessary for the successful application and optimization of extremely thermoacidophilic bioleaching.

2. Biodiversity of Extremely Thermoacidophilic Microorganisms

To date, the only known extreme thermoacidophiles (as defined here, microorganisms with Topt > 65 °C, pHopt < 3.5) belong to the crenarchaeotal class of Thermoprotei, represented by the orders Desulfurococcales, Thermoproteales, Fervidococcales, Acidilobales, and Sulfolobales, with only certain species in the Sulfolobales thus far considered for bioleaching [12,13,14]. The Euryarchaeota order Thermoplasmatales, while containing extremely acidophilic genera, some found in bioleaching, are considered to be moderate thermophiles [15]. The Sulfolobales are comprised of the genera Sulfolobus (9 species), Acidianus (8 species), Metallosphaera (5 species), Sulfurococcus (2 species), Stygioglobus (one species), and Sulfurisphaera (one species). The following section highlights the physiology of the Sulfolobales, with an overview of isolation and sequencing chronology presented in Figure 1 and growth physiology in Table 1.
Table 1. Growth physiology information for the Sulfolobales. (Opt.) Optimum (ED) Electron donor, (EA) Electron acceptor, (Seq.) Genome sequenced, (ND) Not determined, (COR) Complex organic compounds, (AA) Amino Acids.
Table 1. Growth physiology information for the Sulfolobales. (Opt.) Optimum (ED) Electron donor, (EA) Electron acceptor, (Seq.) Genome sequenced, (ND) Not determined, (COR) Complex organic compounds, (AA) Amino Acids.
SpeciesIsolated FrompH (Opt.)T °C (Opt.)EDEAGrowth ModesSeq.Refs.
Sulfolobus acidocaldarius (98-3T, DSM639)Locomotive Spring, Yellowstone National Park, USA1.0–5.9 (2.0–3.0)55–85 (70–75)H2 ~, H2S ?, S0 ?, FeS ?, K2S4O6 ?, COR, Sugars, AAO2Heterotrophic Y[16,17,18,19,20]
Sulfolobus solfataricus (P2, DSM1617)Hot spring, Pisciarelli Solfatara, Italy2.0–4.0 (3.0)65–87 (80)H2 ~, H2S ?, S0 ?, FeS ?, K2S4O6 ?, COR, Sugars, AAO2HeterotrophicY[17,18,19,21,22]
Sulfolobus shibatae (B12T, DSM 5389)Geothermal pool, Beppu, Kiushu Island, JapanND (3.0)ND–86 (81)H2 ~, S0, Sugars, AAO2Heterotrophic MixotrophicN[17,18,23]
Sulfolobus metallicus (Kra23T, DSM 6482)Continental solfataric fields, Iceland1.0–4.5 (ND)50–75 (65)S0, Fe2+, FeS2, CuFeS2, ZnS, CdSO2ChemolithoautotrophicN[18,24,25]
Sulfolobus tokodaii (7T, DSM 16993)Beppu Hot Springs, Kyushu Island, Japan2.0–5.0 (2.5–3.0)70–85 (80)S0, Fe2+, COR, AAO2Heterotrophic MixotrophicY[25,26,27,28]
Sulfolobus yangmingensis (YM1T)Acidic and muddy hot spring, Yang-Ming National Park, Taiwan2.0–6.0 (4.0)65–90 (80)S0, FeS, K2S4O6, COR, Sugars, AAO2Heterotrophic Mixotrophic ChemolithoautotrophicN[29]
Sulfolobus tengchongensis (RT8-4T)Sulfur-rich hot spring, Tengchong, China1.7–6.5 (3.5)65–95 (85)S0, COR, Sugars, AAO2Heterotrophic Mixotrophic ChemolithoautotrophicN[30]
Sulfolobus islandicus (Ren1H1)Solfataric fields, IcelandNDNDNDNDHeterotrophicN ^[31]
Metallosphaera sedula (TH2T, DSM 5348)Thermal pond in Pisciarelli Solfatara, Italy1.0–4.5 (2.0)50–80 (75)H2, S0, K2S4O6, K2SO4, Fe2+, FeS2, CuFeS2, CdS, SnS, ZnS, COR, Sugars, AAO2Heterotrophic Mixotrophic ChemolithoautotrophicY[18,32,33,34]
Metallosphaera prunae (Ron 12/IIT, DSM 10039)Smoldering slag heap, uranium mine, Thüringen, Germany1.0–4.5 (2.0)55–80 (75)H2, So, FeS2, CuFeS2, ZnS, COR, Sugars , AAOHeterotrophic Mixotrophic ChemolithoautotrophicY %[35,36]
Metallosphaera hakonensis (HO1-1T, DSM 7519)Geothermal field, Hakone National Park, Japan1.0–4.0 (3.0)50–80 (70)H2S, S0, K2S4O6, Fe2+, FeS, FeS2, CuFeS2, COR, Sugars, AAO2Heterotrophic Mixotrophic ChemolithoautotrophicN[19,37,38,39,40]
Metallosphaera cuprina (Ar-4T)Sulfuric hot spring in Tengchong, Yunnan, China2.5–5.5 (3.5)55–75 (65)S0, K2S4O6, Fe2+, FeS2, CuFeS2, COR, Sugars, AAO2Heterotrophic Mixotrophic ChemolithoautotrophicY[40,41]
Metallosphaera yellowstonensis (MK1T)Acidic iron mat, Yellowstone National Park, USA1.0–4.5 (2.0–3.0)45–85 (65–75)S0, Fe2+ sorbed, FeS, FeS2, CuFeS2, CuS, ZnS, CORO2Heterotrophic Mixotrophic ChemolithoautotrophicY[42,43]
Acidianus hospitalis (W1)Acidic hot spring, Yellowstone National Park, USA2.0 ?85 ?NDNDNDY[44,45,46]
Candidatus Acidianus copahuensis (ALE1)Copahue geothermal area, Argentina1.0–5.0 (2.5–3.0)55–80 (75)H2,S0, K2S4O6,Fe2+, FeS2, CuS, ZnS, COR, SugarsFe3+, S0, O2Heterotrophic Mixotrophic ChemolithoautotrophicY[47,48]
Acidianus infernus (So4aT, DSM 3191)Solfatara Crater and Pisciarelli Solfatara, Naples1.0–5.5 (2.0)65–96 (90)H2, H2S, S0S0, O2, MO42-Mixotrophic ChemolithoautotrophicN[18,49,50]
Acidianus ambivalens (Lei 10T, DSM 3772)Solfatara, Iceland1.0–3.5 (2.5)70–87 (80)H2, H2S, S0S0, O2Mixotrophic ChemolithoautotrophicN[50,51,52]
Acidianus brierleyi (DSM 1651T)Acid hot spring, Yellowstone National Park1.0–6.0 (1.5–2.0)45–75 (70)H2 ?, H2S, S0, Fe2+,FeS2, CuFeS2, ZnS, MoS2, CORFe3+, S0, O2, MO42-Heterotrophic Mixotrophic ChemolithoautotrophicN[14,18,21,49,50,53,54,55,56]
Acidianus sulfidivorans (JPTT, DSM 18786)Solfatara on Lihir Island, Papua New Guinea0.35–3.0 (0.8–1.4)45–83 (74)H2S, S0 ,Fe2+, FeS2,CuFeSs, FeAsSFe3+, S0, O2Mixotrophic ChemolithoautotrophicN[50]
Acidianus tengchongensis (S5T)Acidothermal spring, Tengchong China1.0–5.5 (1.5–2.0)60–75 (70)H2, S0, S2O32−S0, O2ChemolithoautotrophicN[57]
Acidianus manzaensis (NA-1T)Fumarole in Manza, Japan1.0–5.0 (1.2–1.5)60–90 (80)H2, S0, COR, SugarsFe3+, O2Heterotrophic Mixotrophic ChemolithoautotrophicN[58]
Acidianus manzaensis (YN25)Acidothermal spring, Yunnan China1.0–6.0 (1.5–2.5)50–85 (65)H2, S0, K2S4O6, Fe2+, CuFeS2 ,COR, Sugars, AAS0, O2Heterotrophic MixotrophicN[59]
Sulfurisphaera ohwakuensis (TA-1T, DSM 12421)Acidic hot spring located in Ohwaku Valley, Hakone, Japan1.0–5.0 (2.0)63–92 (84)H2, S0, CORS0, O2Heterotrophic MixotrophicN[60]
Stygiolobus azoricus (FC6T, DSM 6296)Acidic geothermal spring (Furnas Caldeira), São Miguel Island, Azores1.0–5.5 (2.5–3.0)57–89 (80)H2S0Mixotrophic ChemolithoautotrophicN[61]
Sulfurococcus yellowstonensis (Str6karT) Thermal spring, Yellow Stone National Park, USA1.0–5.5 (2.0–2.6)40–80 (60)S0, FeS2, ZnS, CuFeS2, Fe2+, COR, SugarsO2Heterotrophic Mixotrophic ChemolithoautotrophicN[13,62]
Sulfurococcus mirabilis (INMI AT-49T) Crater, Uzon volcano in Kamchatka, Russia1.0–5.8 (2.0–2.6)50–86 (70–75)S0, FeS2, ZnS, CuFeS2, COR, Sugars, AAO2Heterotrophic Mixotrophic ChemolithoautotrophicN[13,62]
Notes: T Type strain; ~ Indicates poor growth; ? Indicates conflicting evidence as growth substrate; ^ Tentative type strain not sequenced but strains sequenced; % Unpublished genome.
Figure 1. Chronology of microbiology advances within the Sulfolobales.
Figure 1. Chronology of microbiology advances within the Sulfolobales.
Minerals 05 00397 g001

2.1. The Genus Sulfolobus

The genus Sulfolobus was defined in 1972 when the first species, Sulfolobus acidocaldarius (98-3), was isolated from Locomotive Spring in Yellowstone National Park, USA [16]. The genus is distributed globally, with species typically isolated from acidic geothermal hot springs. Only seven of the nine species comprising the genus have been characterized in any physiological detail. Sulfolobus species are strict aerobes, with metabolic features ranging from heterotrophy to obligate chemolithoautotrophy. The genus was initially defined by its members’ ability to oxidize elemental sulfur (S0), although subsequent work provided conflicting evidence on this physiological trait.
Heterotrophic growth represents a unifying trait among Sulfolobus species, except for S. metallicus, which is an obligate chemolithoautotroph. The range of substrates supporting heterotrophic growth varies markedly within the genus, though all species utilize complex organic substrates (e.g., yeast extract and tryptone). S. acidocaldarius grows on a limited set of monosaccharides (d-fucose, d-glucose), disaccharides (sucrose), and polysaccharides (maltotriose, dextrin, starch), along with certain amino acids (l-alanine, l-asparagine, l-aspartate, l-glutamate) [17]. S. tengchongensis utilizes more sugars and amino acids than S. acidocaldarius, with pentoses (d-arabinose, d-xylose), hexoses (d-galactose, d-fructose), disaccharides (maltose, sucrose), and amino acids (l-glutamic acid) supporting growth [30]. Compared to S. acidocaldarius, S. solfataricus also grows on a broader range of sugars, including pentoses, hexoses, disaccharides, and polysaccharides [17]. S. shibatae shares many properties with S. solfataricus, but represents a distinct species based on genome content [17,23]. Heterotrophic growth of Sulfolobus tokodaii (7), formerly S. acidocaldarius strain 7, has only been determined for complex organic substrates and certain amino acids, and a complete survey of sugar utilization has not been done [26]. S. yangmingensis is capable of utilizing all sugars observed for other Sulfolobus species, in addition to l-rhamnose, and utilizes all 20 amino acids, except cysteine [29]. None of the tested complex organic substrates, sugars or amino acids supported growth of S. metallicus [24].
The use of inorganic substrates varies widely amongst Sulfolobus species. S. acidocaldarius, S. solfataricus, and S. shibatae can oxidize H2, allowing for mixotrophic growth on H2 and yeast extract, though growth was poor compared to growth of other Sulfolobales [18]. S. metallicus, the only obligate chemolithoautotroph within the genus, cannot grow on H2 as an energy source [18]. H2 utilization by S. tokodaii, S. tengchongensis, and S. yangmingensis was not determined at isolation, nor has it been reported to date [26,29,30]. There is conflicting evidence for the chemolithoautotrophic growth of the type strains of S. acidocaldarius Deutsche Sammlung von Mikroorganismen (DSM) 639 and S. solfataricus DSM 1616. At isolation, both S. acidocaldarius DSM 639 and S. solfataricus DSM 1616 were reported to grow autotrophically on S0, but this observation was contradicted by subsequent work and several key reviews [32,62,63]. The consensus appears to be that these species mutated into obligately heterotrophic strains. However, strain analysis of Metallosphaera hakonensis (formerly S. hakonensis) and S. tengchongensis, which also included S. acidocaldarius DSM 639 and S. solfataricus DSM 1616, showed that these last two Sulfolobus species grew chemolithoautotrophically on S0, FeS, potassium tetrathionate, and H2S [19,30].
S. tokodaii was shown to be incapable of chemolithoautotrophic growth on S0; limited growth was possible on S0 when supplemented with organic carbon [26]. Further work revealed the capacity of S. tokodaii to oxidize Fe(II) with yeast extract supplementation, while weak growth and Fe(II) occurred for autotrophic conditions. [25]. S. tokodaii’s genome encodes genes related to hydrogen sulfide metabolism, though their connection to sulfur metabolism has not been determined. [28]. Chemolithoautotrophic growth of S. yangmingensis and S. tengchongensis by S0 oxidation has been noted; the former utilizes FeS and potassium tetrathionate [29,30].
S. metallicus is distinctive among members of the genus Sulfolobus due to its inability to utilize organic carbon sources for growth, exhibiting obligate chemolithoautotrophy. Inorganic substrates supporting growth include: S0, FeSO4, FeS2, CuFeS2, ZnS, and CdS, but not H2, FeAsS, Cu5FeS4, HgS, Cu2S, CuS, FeS, MoS2, Sb2S3, and SnS [18,24,25]. The ability of S. metallicus to oxidize S0, reduced sulfur compounds, and Fe(II) makes it relevant to heavy metal mobilization applications, particularly bioleaching [64].
S. acidocaldarius and S. solfataricus, S. tengchongensis, and to a lesser extent S. shibatae, appear to be motile [17,23,30], while S. metallicus and S. yangmingensis are immotile [24,29]. It is not clear whether S. tokodaii is motile, but the genome contains the archaellum operon found in S. acidocaldarius, indicating S. tokodaii is likely motile [65]. To date, published genomes exist for S. acidocaldarius (98-3, N8, Ron12/I, SUSAZ), S. solfataricus (P2, 98/2), S. tokodaii (7), Sulfolobus Type II, and a myriad of S. islandicus strains [20,22,28,66,67,68,69,70,71].
S. acidocaldarius, S. solfataricus, and S. islandicus are the only Sulfolobales with genetic tools for molecular biology manipulation [72,73,74,75,76]. In the case of S. acidocaldarius and S. solfataricus, these genetic systems are dependent upon restoration of an engineered auxotrophy. The recent development of a uracil-auxotrophic S. acidocaldarius mutant MW001 has led to a tractable system for metabolic and genetic studies [72], as well as reliable inducible promoters [72,77] and a plasmid-based recombinant expression system [78]. While uracil selection has been utilized in S. solfataricus, more success has been achieved in the utilization of the lactose auxotrophic strain PBL2025 and the lactose/maltose auxotrophic strain PBL2069, which also allow for integration of linear DNA fragments [73,74]. In contrast, S. islandicus faced difficulties in the early stages of developing reliable auxotrophic selection [79]. However, the recent use of simvastatin affords a distinctly different route of selection based on antimicrobials [76,80]. These new genetic systems provide a means to further study and elucidate metabolic and genetic pathways involved in metal mobilization and resistance.
There is limited information on other Sulfolobus species that have been isolated (e.g., S. neozealandicus, [81,82]). S. islandicus strains have been isolated from around the world [31,68,83,84], although the growth physiology of the proposed type strain REN1H1 has not been studied to any significant extent [63]. S. islandicus is a model species for the study of clustered regularly interspaced short palindromic repeats (CRISPR) systems within the Sulfolobales [85,86].

2.2. The Genus Metallosphaera

The type strain of the genus, Metallosphaera sedula (TH2T), was isolated from a thermal pond in Pisciarelli Solfatara (near Naples, Italy) [32]. To date, the genus Metallosphaera includes five reported species with diverse growth physiology. All Metallosphaera species are obligate aerobes, utilizing O2 as their only terminal electron acceptor, and are capable of facultative chemolithoautotrophy, on a variety of inorganic substrates. Variations exists in degrees of heterotrophy in the genus Metallosphaera, with M. cuprina appearing to be the only member capable of significant growth on sugars [40]. M. sedula can grow on beef extract, casamino acids, peptone, trypone and yeast extract, but no utilization of sugars was noted at isolation [32]. However, recent work has indicated M. sedula can use d-mannose and individual amino acids (l-aspartic acid, l-glutamic acid, l-tryptophan, l-alanine), though growth is limited for d-mannose, l-aspartic acid, and l-glutamic acid [40]. Originally, M. prunae was reported to use the same heterotrophic substrates as M. sedula, except for casamino acids and tryptone which were not tested; no utilization of sugars was noted when isolated [35]. Recent work has shown that M. prunae can indeed utilize casamino acids and tryptone, along with d-mannose and individual amino acids (l-aspartic acid, l-glutamic acid, l-tryptophan, L-alanine) [40]. However, growth on d-mannose and l-tryptophan is limited compared to complex organic carbon sources. M. hakonensis exhibits strong heterotrophy on yeast extract, while has limited growth on maltose, l-glutamic acid and l-tryptophan [19]. However, subsequent analysis showed growth on beef extract, peptone, casamino acids, maltose, and l-glutamic acid, with weaker growth achieved on tryptone [40]. M. cuprina differentiates itself from other Metallosphaera species by its broader range of sugar utilization, as it is capable of growth on l-arabinose, d-xylose and d-glucose [40]. M. yellowstonensis growth on sugars and individual amino acids has not been determined, but the organism can grow on YE as the sole carbon and energy source [43].
The unifying trait among Metallosphaera species is their chemolithoautotrophy on S0. H2 oxidation has been determined only for M. sedula and M. prunae [18,35]. Beyond S0 and H2, M. sedula can grow chemolithoautotrophically with inorganic electron donors, including potassium tetrathionate, potassium sulfate, and metal sulfides (FeS2, ZnS, CuFeS2, CdS, SnS) and FeSO4 [18,32,33,87]. However, M. sedula cannot utilize FeAsS, Cu5FeS4, HgS, Cu2S, CuS, PbS, FeS, MoS2 or Sb2S3 as growth substrates [32]. M. prunae is capable of chemolithoautotrophy with inorganic electron donors, including S0, FeS2, CuFeS2, ZnS [35], but thiosulfate, potassium tetrathionate, FeSO4, and various coal substrates do not serve as inorganic energy sources. M. prunae’s reduced capacity to utilize Fe(II) and potassium tetrathionate as an electron donor differentiates it from others in the genus. Growth on H2S for M. sedula and M. prunae has not been reported [40]. M. hakonensis exhibits chemolithoautotrophy, with inorganic electron donors FeS, FeSO4, CuFeS2, potassium tetrathionate, H2S [14,19,37]. Chemolithoautrophic growth for M. cuprina was supported by FeS, potassium tetrathionate, FeSO4, FeS2, CuFeS2 [40]. Unlike M. hakonensis, M. cuprina was not able to utilize H2S as an electron donor. Finally, a detailed analysis of M. yellowstonensis growth physiology revealed inorganic energy sources Fe(II) sorbed to ferrihydrite, FeS, FeS2, CuFeS2, CuS, and ZnS [42]. Growth was not supported by FeCO3, Fe3O4, FeSO4, potassium tetrathionate, or Na2S. Autotrophic growth for M. sedula, like several other Sulfolobales, involves the 3-hydroxypropionate/4-hydroxybutyrate cycle to assimilate CO2 [88]. This pathway is evident in the sequenced genomes of M. cuprina and M. yellowstonensis [41,43].
As indicated above, Fe(II) oxidation is associated with growth for M. sedula, M. cuprina, M. hakonensis and M. yellowstonensis, though for M. yellowstonensis only when Fe(II) is sorbed to ferrihydrite [42]. M. prunae cannot utilize Fe(II) for growth, perhaps because of a mutation [36]. As is the case with S. metallicus, the capability of Metallosphaera species to oxidize S0, reduced sulfur compounds, and Fe(II) make these archaea relevant to heavy metal mobilization, particularly bioleaching [64].
Motility is a shared characteristic for M. cuprina, M. sedula, M. prunae, but not for M. hakonensis; motility has yet to be determined for M. yellowstonensis, though its genome contains the archaellum operon found in S. acidocaldarius, indicating the potential for motility [19,32,35,40,42,65]. To date, M. sedula, M. cuprina, and M. yellowstonensis have published genomes and an unpublished draft genome exists for M. prunae [34,36,41,43]. In fact, the nearly identical genomes of M. sedula and M. prunae suggest that these are strains and not different species [36].

2.3. The Genus Acidianus

Nine species currently comprise the genus Acidianus, all of which are facultative anaerobes capable of chemolithoautotrophy and, in some instances, facultative autotrophy. To date, only the genomes of A. copahuensis and A. hospitalis have been sequenced [46,48]. The common characteristic displayed by all Acidianus species is their ability to oxidize or reduce elemental sulfur, depending on oxygen availability. The only exception is A. manzaensis NA-1, which cannot use S0 as an electron acceptor during anaerobic respiration. Under aerobic conditions, S0 serves as an electron donor and is oxidized to sulfuric acid. While under anaerobic conditions, S0 serves as an electron acceptor and is reduced to H2S. Additionally, all characterized Acidianus species can utilize H2 as an electron donor for aerobic respiration, except A. sulfidivorans.
Under anaerobic conditions, when S0 serves as the electron acceptor, H2 becomes the electron donor. A. infernus, A. ambivalens, A. tengchongensis, A. manzaensis YN25, and A. copahuensis are capable of utilizing H2 as an electron donor, while A. brierleyi and A. sulfidivorans cannot [47,48,50,57,59]. This suggests that A. brierleyi and A. sulfidivorans lack the Ni-Fe-hydrogenase, an essential enzyme used by A. ambivalens for H2 oxidation coupled to S0 reduction [89]. There are conflicting reports as to whether A. brierleyi can utilize H2 as an electron donor for S0 reduction [18,49,50]. Other electron acceptors include Fe(III) coupled with electron donors S0 and H2 for A. copahuensis and A. manzaensis NA-1, and H2S for A. sulfidivorans and A. brierleyi [47,50,58]. Furthermore, A. brierleyi and A. infernus can utilize MO42− as an electron acceptor for S0 oxidation [49].
A. brierleyi, A. copahuensis, A. manzaensis YN25, and A. manzaensis NA-1 are capable of heterotrophic growth, utilizing organic compounds as the sole energy source [47,49,58,59]. The former, along with A. sulfidivorans, are facultative autotrophs, capable of using either organic or inorganic carbon, while A. infernus, A. ambivalens, and A. tengchongensis are obligate autotrophs, solely reliant on inorganic carbon sources. A. copahuensis, A. brierleyi, A. sulfidivorans, and A. manzaensis YN25 can oxidize Fe(II) under aerobic conditions [47,49,50,53,59]. This trait, and the fact that A. copahuensis, A. brierleyi, A. sulfidivorans, and A. manzaensis YN25 utilize various metal sulfides, make them candidates for bioleaching applications [47,50,54,55,56,59].
A. hospitalis growth physiology has not been fully characterized, but genome analysis indicates that it can grow by facultative chemolithoautotrophy. As such, energy sources likely include complex organic compounds, S0, hydrogen sulfide and other reduced inorganic sulphide compounds, but not Fe(II). No Ni–Fe-hydrogenase can be identified in this species, indicating that A. hospitalis does use H2 as an electron donor for growth [46].

2.4. The Genera Sulfurisphaera, Stygiolobus and Sulfurococcus

The only current member of the genus Sulfurisphaera, S. ohwakuensis (TA-I), was isolated from an acidic hot spring located in Ohwaku Valley, Hakone, Japan [60]. S. ohwakuensis is a facultative anaerobe, utilizing S0 and O2 as electron acceptors, and H2 along with complex organic compounds yeast extract and peptone as electron donors. The organism does not exhibit autotrophy, nor were pili- or flagella-like structures found.
Stygiolobus azoricus (FC6) was isolated from an acidic geothermal spring on São Miguel Island, Azores [61]. St. azoricus differentiates itself from all other Sulfolobales in being strictly anaerobic, but shares with Acidianus spp. the ability to grow chemolithoautotrophically by S0 using H2 [61]. The microorganism displays no flagella or motility, but is surrounded by pilus- or fimbria-like appendages.
The genus Sulfurococcus is represented by two species S. yellowstonensis and S. mirabilis, both facultative chemolithoautotrophs [13,62]. Both species use complex organic compounds and sugars for growth, but S. mirabilis can also utilize amino acids. While these Sulfurococcus species use S0, FeS2, ZnS, CuFeS2, S. yellowstonensis also uses Fe(II). S. yellowstonensis capacity for growth on sulfur, mineral sulfides and Fe(II) makes it relevant to bioleaching, as is the case for other Sulfolobales noted above.

2.5. Sequenced and Unclassified Sulfolobales

One novel Sulfolobales archaeon, named Acd1, was isolated during a metagenomic study of nanoarchaeon from Obsidian Pool, Yellowstone National Park and has an available genome sequence [90]. Another, Sulfolobales archaeon AZ1 was isolated from a hot spring located at Los Azufres National Park, Mexico and has been proposed as Candidatus Aramenus sulfurataquae, representing a novel genus and species [91].

3. Biooxidation of Heavy Metals

Pioneering work by Ingledew and co-workers laid the framework for studies on microbial Fe(II) oxidation by the mesophilic bacterium Acidithiobacillus ferrooxidans, thereby forming a foundational basis to study Fe(II) oxidation pathways [15,92,93,94,95,96]. Extensively characterized, A. ferrooxidans has been the main microorganism considered for biohydrometallurical processes, but challenges with the recalcitrant nature of ores like chalcopyrite motivates the search for other bioleachers. Understanding the basic elements of energy metabolism in heavy metal mobilizing microorganisms is critical for future technological applications, especially when solutions tailored to specific ores are needed.
Initial efforts to understand the mechanisms driving the oxidation of metals by the Sulfolobales began more than 20 years ago, although it was only recently that significant progress has been reported in this regard. From the beginning, the respiratory clusters associated with Fe(II) oxidation of the extremely thermoacidophilic Sulfolobales could be differentiated from their mesophilic counterparts. Fe(II)-grown cells of M. sedula and A. brierleyi showed high expression of a novel membrane-bound yellow cytochrome, directly reduced by Fe(II), possibly representing a unique extremely thermoacidophilic redox-active enzyme associated with respiratory Fe(II) oxidation [97]. Sulfolobus strain BC, now S. metallicus, produced copious amounts of a similar novel acid-stable material during growth on Fe(II), revealing similarity among the Fe(II)-oxidizing archaea [97,98]. The results suggested phylogenetically distinct groups of Fe(II)-oxidizing organisms have characteristically unique acid-stable, redox-active biomolecule.
Until the early 2000s, most studies on crenarchaeotal respiratory chains focused on understanding the molecular properties of oxidases and associated electron transfer proteins [99,100,101]. The SoxABCD-SoxL complex, an aa3 quinol oxidase [102,103,104], the SoxM supercomplex, a bb3 terminal oxidase [105,106,107,108], and the CbsAB-SoxLN complex, a cytochrome ba [109] of S. acidocaldarius, later found in A. ambivalens [110], and the DoxBCE complex, an aa3-type quinol oxidase [111,112,113] of A. ambivalens had encapsulated the current view of aerobic respiratory electron transfer in the Sulfolobales. Unfortunately, neither S. acidocaldarius nor A. ambivalens can grow well on metal sulfides, thus motivating study of Crenarchaeota capable of growth on metal sulfides.
M. sedula, capable of chemolithoautotrophy with metal sulfides (e.g., pyrite) or sulfur, and heterotrophy with complex organic substrates, is a prime candidate for investigation of Fe(II) oxidation mechanisms within the Sulfolobales. Prior to the availability of the M. sedula genome sequence, three gene clusters containing oxidases and cytochromes were observed to be differentially expressed, according to whether growth was chemolithoautotrophic (S0 or FeS2) or heterotrophic (yeast extract). M. sedula’s homologs to soxB, representative of the cluster, and soxM were highly expressed for growth on S0 and yeast extract, respectively [114]. The soxL2N transcriptional unit, separately located from csbA, exhibited high expression on either S0 or yeast extract. Growth on S0 and FeS2 induced csbA, such that it was the highest transcribed gene for FeS2, indicating the gene product’s importance for chemolithoautotrophic growth. CsbA is a membrane-bound cytochrome b566/588, implicated in electron shuttling across the pseudo-periplasmic space of S. acidocaldarius and speculated to be related to the previously mentioned novel yellow redox-active enzyme [97,114,115,116]. While a potential chemolithoautotrophic electron shuttle had been identified, the corresponding Fe(II)- and/or sulfur-oxidizing enzymes remained uncharacterized.
Analysis of S. metallicus and S. tokodaii grown on Fe(II) finally revealed a suspected genetic basis for Fe(II) oxidation among the Sulfolobales. The ferrous iron oxidation (fox) genes, encoding a novel terminal oxidase complex first characterized in S. metallicus, were highly induced during Fe(II) oxidation, with homologs present in S. tokodaii, capable of Fe(II) oxidation, but not S. acidocaldarius, incapable of Fe(II) oxidation [25]. The significant involvement of the fox genes in Fe(II) oxidation was further supported by the observation that pyrite-grown cells, but not sulfur-grown cells, exhibited a dominant membrane-bound protein corresponding to FoxA. Additionally, the observations suggest that one of the fox genes is a more likely candidate, than csbA, for the previously noted redox-active enzyme, associated with Fe(II) oxidizing Sulfolobales [25,97,98,114]. The availability of genome sequence data for the Sulfolobales has established the presence of the fox gene cluster in Fe(II)-oxidizing species, and the absence of this cluster in non-Fe(II)-oxidizing species, except for M. cuprina (Table 2). The genome of M. prunae contains the fox cluster, but with a mutated foxA’, possibly impacting activity of the cluster as indicated by a reduced capacity for uranium and Fe(II) oxidation [36]. The presence of the fox cluster appears to correlate to Fe(II)-oxidizing capacity. However, the fact that the M. cuprina appears not to encode this cluster, but oxidizes Fe(II), needs to be resolved.
Results from transcriptomic studies continue to support the hypothesis of significant involvement of the fox gene cluster in Fe(II) oxidation, though in the absence of biochemical characterization other candidates cannot be completely ruled out [33,43]. As previously noted, the cbsA-soxLN2 complex exhibited high expression levels in M. sedula when grown on Fe(II) and S0, with further work revealing preferential differential up-regulation for Fe(II) [33,114]. In contrast, analysis of M. yellowstonensis showed that cbsA expression was the same in the presence and absence of Fe(II) [43]. Furthermore, csbA-soxNL2 homologs are present in non-Fe(II)-oxidizing Sulfolobus species. In light of the previous observations, the cluster cannot be ruled out for Fe(II) oxidation.
Other likely Fe(II) oxidation candidates include the other three terminal oxidase clusters, namely SoxABCD, SoxM supercomplex and DoxBCE. However, gene expression and physiological analysis point to functions of sulfur oxidation for SoxABCD and DoxBCE, and heterotrophy for the SoxM super-complex [33,34,43,114]. As with csbA-soxNL2, homologs are present in non-Fe(II)-oxidizing Sulfolobus species. The function of rusticyanin, sulfocyanin, and other novel multi-copper oxidases has yet to be determined for extremely thermoacidophilic Fe(II)-oxidizers, though they have been hypothesized to function as electron shuttles to the terminal oxidase. Rus-like blue copper proteins in Metallosphaera spp., with plastocyanin type I domain, were highly transcribed on growth on chalcopyrite and triuranium octaoxide, begging the question about the cellular localization of Rus and its role in the electron transport chain for Fe(II) oxidation in Metallosphaera spp. [33,36].
Table 2. Complete terminal oxidase clusters and associated proteins identifiable in sequenced extreme thermoacidophile genomes. (x) and (-) indicate presence and absence, respectively, with (y) being used when a secondary candidate exists. Species in bold are known to oxidize Fe(II). Blank boxes indicate absence of sequence data for those species. Homology searches were completed using previously identified M. sedula proteins as search queries with The National Center for Biotechnology Information’s (NCBI) Basic Local Alignment Search Tool (BLAST) against the Sulfolobales (taxid:2281) database.
Table 2. Complete terminal oxidase clusters and associated proteins identifiable in sequenced extreme thermoacidophile genomes. (x) and (-) indicate presence and absence, respectively, with (y) being used when a secondary candidate exists. Species in bold are known to oxidize Fe(II). Blank boxes indicate absence of sequence data for those species. Homology searches were completed using previously identified M. sedula proteins as search queries with The National Center for Biotechnology Information’s (NCBI) Basic Local Alignment Search Tool (BLAST) against the Sulfolobales (taxid:2281) database.
Terminal OxidaseFox ABCDEFGSox ABCDLSox EFGHIMDoxBCECbsAB-SoxL2NRusticyaninSulfocyanin
Metallosphaera sedulaxxxxxx, yx, y
Metallosphaera prunaexxxxxx, yx, y
Metallosphaera cuprina-xxxx-x, y
Metallosphaera yellowstonensisxxxxxx, yx, y
Sulfolobus solfataricus-xxxxxx, y
Sulfolobus acidocaldarius-xxxx-x, y
Sulfolobus tokodaiixxxxx-x, y
Sulfolobus metallicusx
Sulfolobus islandicus-xxxxx, yx, y
Acidianus brierleyi x
Acidianus ambivalens xx
Acidianus hospitalis---xx-x
Candidatus Acidianus copahuensisxx-xxxx, y
Notes: Gene IDs used for BLAST queries: FoxA (Msed_0484), FoxB (Msed_0480), FoxC (Msed_0478), FoxD (Msed_0477), FoxE (Msed_0475), FoxF (Msed_0474), FoxD (Msed_0469); SoxA (Msed_0289), SoxB (Msed0290), SoxC (Msed_0288), SoxD (Msed_0285), SoxL (Msed_0287); SoxE (Msed_0323), SoxF (Msed_0322), SoxG (Msed_0321), SoxH (Msed_0320), SoxI (Msed_0219), SoxM (Msed_0324); DoxB (Msed_2032), DoxC (Msed_2031), DoxE (Msed_2030); CbsA (Msed_0504), CbsB (Msed_0503), SoxL2 (Msed_0501), SoxN (Msed_0500); Rusticyanin 1 (Msed_0966), Rusticyanin 2 (Msed_1206); Sulfocyanin 1 (Msed_0323), Sulfocyanin 2 (Msed_0826).
There are additional membrane bound redox complexes which may respond to different organic and inorganic substrates: Nicotinamide adenine dinucleotide reduced (NADH):quinone oxidoreductase from A. ambivalens and S. metallicus is proposed to transfers electron from NADH to quinones [117], while the succinate:quinone oxidoreductase SdhABCD from A. ambivalens and S. tokodaii is proposed to transfer electrons from succinate to quinones [118,119]. The thiosulfate:quinone oxidoreductase, tetrathionate hydrolase TetH from A. ferrooxidans and Acidithiobacillus caldus [120,121] functions in S0 oxidation, in agreement with the M. sedula model [33,96]. The DoxDA, aa3 type quinol oxidase, has been annotated as a thiosulfate:quinone oxidoreducatase (TQO) in A. ambivalens [111], with likely involvement in sulfur oxidation. It has been shown that TQO can oxidize thiosulphate to tetrathionate, using ferricyanide or decyl ubiquinone (DQ) as electron acceptors [122].
As noted above, biooxidation of Fe(II) has been most widely and extensively studied in A. ferrooxidans and a brief overview of the current model is warranted prior to discussion of models within the Sulfolobales. The overall organization of Fe(II) oxidation components, mainly the vertical topography where Fe(II) is kept outside the cell, appears to be conserved, but significant diversity exists among the redox proteins [15]. For A. ferrooxidans, the transfer of electron from Fe(II) to oxygen involves a super-complex connecting the outer and inner membranes. The super-complex consists of an outer membrane high molecular-weight cytochrome c, encoded by cyc2, where Fe(II) oxidation occurs [123,124], a gene of unknown function (ORF1), a periplasmic soluble blue copper protein rusticyanin encoded by rus believed to responsible of uphill/downhill bifurcation [125,126], and a periplasmic membrane-bound di-hemic cytochrome c4 encoded by cyc1 [127]. Downhill flow proceeds to a terminal aa3-type cytochrome oxidase encoded by the coxBACD gene cluster [128,129]. The uphill components flow proceeds to a cytochrome bc1 complex (complex III, ubiquinol-cytochrome c reductase) through the quinone pool, and finally to a NADH1 dehydrogenase complex [130,131,132]. The bc1 complex is part of a five-gene operon, termed the petI operon, which is adjacent to the resBC operon, suspected to be involved in the construction of the c1 cytochrome [96]. Elements remaining to be determined in the electron transport chain are the specific interactions between certain complexes, assembly proteins, and the mechanisms of regulation for modulating uphill or downhill flux [96].
To date, hypothetical models for Fe(II) oxidation by the Fox cluster has been proposed for Metallosphaera species, based on expression, modeling and comparative genomic analysis (Figure 2) [33,43]. The Sulfolobales do not have the initial electron acceptor from Fe(II) found in A. ferrooxidans, a c-type cytochrome, supporting the notion that the Sulfolobales Fe(II) oxidation pathway is evolutionarily distinct [15]. Initially, electrons are extracted from Fe(II) by FoxCD, cytochrome b, and shuttled by a multi-copper oxidase either uphill or downhill. FoxA1 and FoxA2 are annotated as cytochrome c-oxidases (subunit I), forming a complex with FoxB, annotated as cytochrome c-oxidase (subunit II), and receive electrons proceeding in the downhill direction from the multi-copper oxidase. FoxG has been annotated as a 4Fe–4S polyferredoxin-like protein, and can form a complex with FoxCD for uphill electron flow through the multi-copper oxidase to the cytochrome ba complex, CbsAB-SoxLN. Electrons then pass through the quinone pool, finally to a NADH dehydrogenase. FoxH has been annotated as a signal transduction protein and its location in the fox cluster suggests an Fe(II)-sensing role.
Recently, M. sedula was found to oxidize uranium trioxide, with the fox cluster likely mediating the oxidation process [36]. The conclusion was supported by M. prunae’s inability to transform the oxide and a corresponding frame shift in foxA’, possibly also exerting a polar effect on the fox cluster. Prior to this report, three microorganisms possessed the ability to oxidize tetravalent U(IV) to hexavalent U(VI), namely the aerobic acidophilic chemolithotroph A. ferrooxidans, previously discussed, the anaerobic chemoorganoheterotroph Geobacter metallireducens, and the anaerobic obligate chemolithotroph Thiobacillus denitrificans [133,134,135]. The latter two catalyze the nitrate-dependent oxidation of U(IV). Two di-heme, c-type cytochromes, putatively c4 and c5 cytochromes, have been found to play a major role in the nitrate-dependent U(IV) oxidation by T. denitrificans [136]. The two cytochromes are membrane-associated and may be periplasmic, based on homology to characterized c4 and c5 cytochromes in Pseudomonas stutzeri. The fact that periplasmic, rather than outer membrane, proteins are involved in the oxidation of UO2 suggests that U(IV) dissolution occurs before U(IV) oxidation, because it is unknown how periplasmic proteins would interact with a solid mineral substrate. Siderophores could enhance the solubility of U(IV), making it more bioavailable to the periplasmic cytochromes, or perhaps some yet undetermined outer membrane protein directly contacts the UO2. The biological oxidation of uranous sulfate, a soluble U(IV) species, by A. ferrooxidans has been demonstrated [133]. The authors hypothesized that rusticyanin was the first protein in the electron transport chain for the uranous ion. Subsequent electron transfer involved a yet unidentified electron acceptor between rusticyanin and cytochrome c. Based on more recent evidence for Fe(II) oxidation, the initial electron acceptor could be Cyc2, as opposed to rusticyanin. This is supported by the fact that the uranous ion has been found to be a competitive inhibitor of Fe(II) oxidation, which would implicate use of the same cytochrome c [137].
Figure 2. Proposed model for Fe(II) oxidation in M. sedula, based on transcriptional response experiments and bioinformatics analysis [33,43]. FoxC is believed to be the primary electron acceptor from metal ions and transfer the electrons to a blue copper protein (Rus—rusticyanin), which can follow an uphill electron flow to NADP+ or a downhill electron flow to O2. The dotted line shows the direction of electron flow. (a) heme a center in FoxA, (CuBa3) binuclear center in FoxA where O2 is reduced to H2O; (b) heme b in FoxC and soxN, (CuA) copper center in FoxB, (Fe–S) iron sulfur clusters in FoxG and SoxL, (Q) ubiquinone, (QH2) hydroquinone.
Figure 2. Proposed model for Fe(II) oxidation in M. sedula, based on transcriptional response experiments and bioinformatics analysis [33,43]. FoxC is believed to be the primary electron acceptor from metal ions and transfer the electrons to a blue copper protein (Rus—rusticyanin), which can follow an uphill electron flow to NADP+ or a downhill electron flow to O2. The dotted line shows the direction of electron flow. (a) heme a center in FoxA, (CuBa3) binuclear center in FoxA where O2 is reduced to H2O; (b) heme b in FoxC and soxN, (CuA) copper center in FoxB, (Fe–S) iron sulfur clusters in FoxG and SoxL, (Q) ubiquinone, (QH2) hydroquinone.
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4. Heavy Metal Resistance Systems in Extreme Thermoacidophiles

The dichotomy between metabolic requirements for metals by microorganisms and the potential associated toxicity has created an interesting and broad set of metal homeostasis and resistance systems required to maintain a delicate balance [138]. The concern of metal toxicity is particularly pertinent to metal mobilizing microbes, which must resist toxic heavy metals released into the environment as a result of their energy metabolism. In general, there are seven broadly defined categories of postulated mechanisms for metal resistance/tolerance in microorganisms (Figure 3): (1) passive tolerance; (2) metal exclusion by permeability barrier; (3) active transport of the metal; (4) intracellular sequestration of the metal by protein/chelator binding; (5) extracellular sequestration of the metal by protein/chelator binding; (6) enzymatic detoxification of the metal to a less toxic form, and (7) reduction in metal sensitivity of cellular targets to metal ions [139]. Microorganisms may contain one or more combinations of the above resistance mechanisms, but the primary mechanism for regulating intracellular metal concentrations under normal growth conditions involves membrane transport. However, exposure to higher toxic concentrations can elicit other more stringent responses that reduce non-specific uptake or induce specific metal resistance mechanisms, for example efflux [140]. The following section describes the current knowledge of metal resistance mechanisms with an emphasis on extremely thermoacidophilic microorganisms, useful for bioleaching applications. Comprehensive reviews covering microbial metal resistance exist, but few specifically target extreme thermoacidophiles [141,142,143,144,145,146].
Figure 3. Overview of metal resistance mechanisms for acidophiles. Energy for transporters can be provided by ATP (P-type ATPase), proton gradient (RND), or chemiosmotic (CDF). Metal sequestration can occur through small molecule complexing agents (e.g., phosphate) or metal-chelating proteins. The exterior blue barrier represents some external permeability barrier (e.g., S-layer or biofilm). The figure does not including reduction in metal sensitivity of cellular targets.
Figure 3. Overview of metal resistance mechanisms for acidophiles. Energy for transporters can be provided by ATP (P-type ATPase), proton gradient (RND), or chemiosmotic (CDF). Metal sequestration can occur through small molecule complexing agents (e.g., phosphate) or metal-chelating proteins. The exterior blue barrier represents some external permeability barrier (e.g., S-layer or biofilm). The figure does not including reduction in metal sensitivity of cellular targets.
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4.1. Passive Tolerance and Metal Exclusion

Recent literature concerning the high capacity of acidophiles to tolerate significantly higher levels of metal ions than their neutrophilic counterparts has revealed a potentially important set of passive tolerance features [141,147]. The first passive mechanism relates to metal ion chelation by sulfate ions, which are typically high in acidophile habitats and are coupled to metal concentrations. Chelation significantly reduces the availability of free ions, which are much more toxic [147,148]. However, soluble complexes can still exert significant toxicity, as found for zinc phosphate with the neutrophile Arthrobacter sp. [149]. Unlike neutrophiles, acidophiles maintain an inside positive cytoplasmic transmembrane potential, thus generating a chemiosmotic gradient inhibiting proton and metal cation passage across the membrane [147,150,151]. At lower pH, a greater competition exists between protons and metal cations for metal-binding sites [152,153], which has been hypothesized to account for decreased toxicity of zinc at lower pH values for acidophiles [154]. However, for more toxic metals, the importance of the above passive systems might be limited [147]. For example, the capacity of an M. sedula strain, deficient in an active efflux system for copper, failed to mobilize chalcopyrite, despite the concomitant increase of sulfate during mineral dissolution [155].
Metal exclusion represents another general defense against toxic metal effects and involves alterations in the cell wall, membrane, envelope, or surface layer (S-layer) in an attempt to prevent damage to intracellular or cytoplasmic targets [139]. Arsenate can be taken up by phosphate transport systems in bacteria, but enhanced resistance can be achieved with highly specific phosphate transporters, excluding arsenate. Specifically, for E. coli, this involves use of the Pst system as opposed to the less specific Pit system [156]. A loss of function for the Pit system in M. sedula increased resistance compared to a spontaneous mutant harboring a restored version, but found to reduce resistance to copper [157]. The result is consistent with mutations of low-affinity, high velocity transporters pit and corA being more tolerant to arsenate and cobalt, respectively [158,159,160,161]. Therefore, a potential system of defense relies on the mutation of non-specific systems or use of more specific uptake systems for essential nutrients in an attempt to exclude toxic metals and avoid the “open gate” issue [140,162]. Limitations exist for this strategy, as use of more specific uptake systems generate tolerant mutants that are less robust than the wild-type [144]. Another resistance strategy exists for metals, such as nickel and cobalt, where repression of permeases responsible for their uptake can prevent associated toxic effects when extracellular concentrations become physiologically dangerous [163].
Non-specific binding of metals by the outer membrane, envelope, S-layer, extracellular polymeric substances (EPS), and/or lipopolysaccharide (LPS) offers yet another means of metal exclusion. Biofilms, generally composed of extracellular polymeric substances, are capable of enhancing metal tolerance of attached communities since they sorb metals [164,165,166]. The extracellular matrix and S-layer, known to contain many functional groups capable of interacting and trapping metals, have in the case of Bacillus sphaericus been shown to act as a protective uranium immobilizing matrix, resulting in a local detoxification [167,168,169,170]. This offers limited metal protection due to saturation of binding sites, but regeneration of binding sites by enzymes, such as phosphatases, can extend the effectiveness of the system [169].

4.2. Copper

Among the metals studied in extreme thermoacidophiles, copper has received the most attention because of the potential of these archaea for extraction of copper from primary ores like chalcopyrite. The known copper resistance strategies employed by these microorganisms includes active transport and metal sequestration. The active transport system utilizes members of the P-type ATPase superfamily, which includes many members responsible for pumping cations against steep electrochemical gradients by exploiting the energy from ATP hydrolysis [171]. A myriad of microorganisms, including the widely studied S. solfataricus, employ single-unit membrane class IB heavy metal translocating P-type ATPases [171,172,173,174], used for metal (Cu2+, Cu+, Ag+, Pb2+, Zn2+, Cd2+, Co2+) homeostasis and resistance [175,176,177,178]. The metal sequestration system is associated with inorganic phosphate metabolism, known to be involved in bacterial stress responses [179,180,181,182]. The role of polyP metabolism in metal stress response among extremophiles has received limited attention, despite the potential of this system to confer significantly higher levels of copper resistance [183,184].
The S. solfataricus genome exhibits only two P-type ATPases, both belonging to class 1B, CopA and CopB encoded by Sso_2651 and Sso_2896, respectively [22]. CopA and CopB are known to impart copper resistance to S. solfataricus, with CopA being an effective copper pump at low and high copper concentrations, and CopB apparently functioning as a low-affinity copper export ATPase extending resistance at higher concentrations [178]. Similar roles have been observed for the homologous CopA and CopB of Thermus thermophilus, with stimulation of each occurring most in the presence of Cu(I) and Cu(II), respectively [185]. Interestingly, PIB-type copper pumps of the CopA-2 subclass have been implicated in the assembly of metalloproteins, such as the copper-containing cbb3-type respiratory oxygen reductases [186,187,188]. As a result, CopA and/or CopB may function together with periplasmic copper chaperons in the assembly of the ba3-type and caa3-type copper-containing respiratory oxygen reductases present in T. thermophilus [185]. However, Völlmecke et al. (2012) demonstrated neither PIB-type ATPase, both of the CopA-1 subclass, found in S. solfataricus play an essential role in cytochrome oxidase biosynthesis.
The CopA operon occurs as the gene cluster copRTA, encoding the copper-responsive regulator CopR [189], the copper-binding protein CopT containing the metal coordinating ligands within the so-called trafficking, resistance and sensing of heavy metals (TRASH) domain [190], and the Cu(I) transporting P1B-type ATPase, which are induced under the presence of excess copper and represent the general structure of the operon found in archaea [191,192]. The copB gene cluster is organized in a different region, with the transcriptional regulator copY and a small copper chaperone of the heavy metal associated (HMA) group copZ arranged in the opposite orientation of the Cu(II) transporting P1B-type ATPase, copYZ/copB [178]. The catalytic ATP-binding/phosphorylation domain of CopB was shown to be active in the presence of Cu(II), but not Cu(I), and is believed to play a role in the transport of Cu(II) [193]. In the hyperthermophilic, sulfate-reducing archaeon Archaeoglobus fulgidus, two copper-transporting ATPases, CopA and CopB, were isolated, characterized and found to activated on Cu(I) and Cu(II), respectively [194,195]. The CopA of A. fulgidus has been extensively studied with regards to its metal binding, actuator, ATP binding domains, and interaction with chaperones [196,197,198,199,200,201,202].
The genome of the extreme acidophilic archaeon Ferroplasma acidarmanus contains a cop operon encoding a putative transcriptional regulator (copY), a putative metal-binding chaperone (copZ), and a putative copper-transporting P-type ATPase (copB) [190]. Transcript levels of the co-transcribed copZB were found to increase in response to exposure to high levels of Cu2+ [203]. Recently, a similar copA operon to the one in S. solfatariucus was studied in M. sedula, which has 20 times greater resistance to Cu2+ [32,34,155,189]. A genetics-based investigation proved the functional role of the PIB-type ATPase operon copRTA found in the M. sedula genome [155]. Further, targeted recombination of copA compromised both metal resistance and eliminated chalcopyrite bioleaching. Two non-identical cop loci in S. metallicus have been identified to respond to both copper and cadmium, implicating functionality in resistance response [204].
Despite the possibility of polyP to impart broad metal resistance, study of polyP related to stress responses has largely been ignored in extremophiles, especially those best suited for biomining applications, except for the bacterium A. ferrooxidans and the archaeon S. metallicus [182,205,206]. Several acidophilic organisms, including A. ferrooxidans, S. metallicus, S. acidocaldarius, A. thiooxidans, M. sedula, A. caldus and to a lesser extent S. solfataricus, accumulate polyP [146,183,184]. Comparison of polyP production in S. solfataricus to that in S. metallicus showed that S. metallicus had significantly higher polyP synthesis and could tolerate up to 200 mM copper sulfate, while S. solfataricus could not resist more than 1–5 mM copper sulfate, suggesting a relationship between Cu-resistance and polyP levels [184]. Also, a study on the transcriptional and functional genes related to survival in the presence of copper for A. ferrooxidans identified polyP as contributing to copper resistance [207]. Another system found in A. ferrooxidans, but not found in the Sulfolobales, is the proton-driven Cus CBA-transport system, studied extensively in E. coli [146,208]. Both the Cop and Cus mechanisms are believed to be key determinants in the copper resistance of A. ferrooxidans [207].
Enzymes essential to polyP metabolism are the polyphosphate kinase (PPK) that catalyzes the reversible conversion of ATP’s gamma phosphate into polyP and the exopolyphosphatase (PPX) that hydrolyzes terminal residues of polyP [206]. Interestingly, production of polyP has been proven to occur in several archaeal species, but only PPX proteins and genes have been described, in particular for S. solfataricus [209]. Further, analysis of extremophilic archaea has shown that Crenarchaeota possess ppx genes but lack ppk genes, while Euryarchaeota possess ppk genes, but no ppx genes. Most extremophilic bacteria included in the analysis contained both genes [206]. The role of polyP in metal resistance in extremophiles encompasses both its role as an energy source and metal chelating agent. Through the action of PPX, the microorganism can generate organic phosphate for metal chelation, or the reversible reaction catalyzed by PPK can generate additional ATP for heavy-metal efflux systems or other cellular metabolism associated with metal challenge [142]. The importance of phosphate metabolism, specifically import via an archaeal Pit system, to enhanced copper resistance for an M. sedula mutant has been established and indicates phosphate plays a key role in supernormal copper resistance [157].

4.3. Mercury

Currently, the most pervasive and generally employed mercury resistance strategy among bacteria and archaea occurs through volatilization [210,211,212,213,214]. Mercury methylation or reduction can lead to volatilization, but only the latter is believed to operate as a resistance mechanism, because organomercurials are highly toxic [144]. Bacterial mediated mercury methylation occurs anaerobically, is coupled to dissimilatory reduction of various electron acceptors, mostly sulfate [215], and has been extensively studied and recently reviewed [216,217]. There exists a limited understanding of mercury methylation in archaea, but it has been shown to occur in certain methanogens [218]. The known mercury resistance strategy in extreme thermoacidophiles is based on reduction of Hg2+ to the volatile Hg0+ by the Hg-reductase MerA and is a homolog of the thoroughly investigated bacterial mercury resistance system (mer) encoded by the mer operon [144,210,213,219,220]. The extreme thermoacidophile mechanism is based on genes encoded by the merRAHI operon, which has been studied in detail for S. solfataricus [221,222]. The gene merA encodes a protein that is homologous to the bacterial mercury reductase MerA. MerR acts as a negative regulator inducing transcription without leaving the merA promoter. MerH contains the conserved metal binding TRASH domain and is suspected to chaperone mercury for mobilization. MerI has a yet undetermined functional role [221,222]. Like other early evolving microbial lineages, Aquifica [223] and Thermus/Deinococcus [224], S. solfataricus’s (Crenarchaeota) mer operon encodes fewer functional genes than the operons found in Proteobacteria, Firmicutes, and Actinobacteria [211]. Another avenue for mercury reduction beyond the mer system has been discovered in A. ferrooxidans, where a cytochrome c oxidase was found to detoxify mercurial compounds [225,226].

4.4. Other Metals (Arsenic, Cadmium, Nickel, Uranium)

Although most work to date centers on copper and mercury resistance, other metals have been studied. Arsenic resistance systems include arsenate reduction followed by arsenite efflux, complexation by metallothioneins, and methylation [227,228,229]. The most pervasive system employs an arsenic resistance (ars) operon encoding an As3+ responsive transcriptional repressor (ArsR) [230], an arsenate reductase (ArsC) responsible for extending resistance to As5+ by mediating As(V) reduction to As(III) [231], which is then extruded by the ArsB antiporter, catalyzing exchange of As(OH)3 for protons and thus conferring resistance [232]. Additionally, some ars operons contain an ArsB complexing As3+-translocating ATPase (ArsA), enhancing resistance [233], and an arsenic metallochaperone (ArsD) that transfers As3+ to ArsA, increasing its ability to extrude arsenite [234,235]. Other genes associated with ars operons include the putative thioredoxin reductase (arsT) [236,237] or thioredoxin system (arsTX) [238] required for As(V) reduction using NADPH reducing power, and two genes of unknown function with weak homology to oxidoreductases (arsO and arsH) [236,239,240]. The ArsH protein from Shigella flexneri was shown to have NADPH-dependent FMN reductase activity [241].
Several archaeal genomes contain homologs of the ArsC [162], but a sub-section that contain other ars operon components, or are known to be resistant to arsenate, appear to lack ArsC. The extreme arsenite resistance of F. acidarmanus has been attributed to the established ars operon, which for this organism is only represented by the arsenite inducible operon homologous to arsRB [242]. Despite the absence of a homolog to the arsenate reductase (arsC), the inability to reduce arsenate and accumulation of intracellular arsenate, F. acidarmanus still possesses extreme arsenate resistance [242,243]. Since ArsB can only extrude As(III), an unknown and novel arsenate resistance mechanism likely exists in F. acidarmanus, with modes including direct efflux of As(V), intracellular sequestration, resistance of cellular components to As(V), or high levels of intracellular phosphate [242]. F. acidarmanus genome does contain a homolog to arsA, but due to the lack of a promoter and an N-terminal domain it is likely a pseudogene [242]. A similar arrangement can be found in other extreme acidophilic archaea, such as Thermoplasma acidophilum and Picrophilus torridus, with the addition of either a partial or complete separately encoded arsA, respectively [244,245]. The genomes of both S. solfataricus and M. sedula encode a stand-alone arsenite transporter ArsB [34]. Transcriptional analysis of the archaea Pyrobaculum aerophilum utilizing arsenate as the terminal electron acceptor revealed the up-regulation of a putative arsR homolog, but not the up-regulation of an annotated arsenical pump-driving ATPase and arsenite permease [246]. In contrast to acidophilic archaea, arsC containing operons are present in other acidophilic bacteria, such as Acidithiobacillus ferrooxidans [239], Acidithiobacillus caldus [247], Acidiphilium multivorum [248], and Leptospirillium ferriphilum [249].
Beyond F. acidarmanus, ars-based arsenic resistance in archaea has only been extensively characterized in Halobacterium sp. strain NRC-1. The megaplasmid pNRC100 encodes the gene clusters arsADRC and arsR2M, while arsB occurs on the chromosome [250]. Deletion of the arsADRC cluster resulted in increased sensitivity to arsenite and antimonite, while deletion of arsB caused no change in sensitivity to either arsenate or arsenite, indicating Halobacterium sp. strain NRC-1 contains a novel arsenite/antimonite extrusion system vastly different from bacterial counterparts [250]. The arsM gene was determined to be a putative methyltransferase, known to exist in mammals, and knockout of the gene produced sensitivity to arsenite, possibly indicating a novel detoxification stragtegy [250]. Analysis of microbial genomes identified 125 bacterial and 16 archaeal homologs of arsM genes, with a subset located downstream of an arsR gene, suggesting these ArsMs confer arsenic resistance [251]. The system now represents an established arsenic resistance system for certain archaea and bacteria [162].
Arsenite oxidation could represent an alternative or enhancing strategy to other known arsenic resistance systems [227,252]. The membrane fraction S. metallicus (formerly Sulfolobus acidocaldarius strain BC) has been shown to oxidize arsenite to the less toxic arsenate using an unknown oxidase [253]. Recently, A. brierleyi was shown to oxidize arsenite from refinery wastewater by an undetermined mechanism, presumably an arsenite oxidase [254]. During the bioleaching of arsenic containing ores and concentrates, considerable care must be exercised as mineral dissolution releases arsenite and unless sufficient Fe(III) is present to oxidize As(III), toxicity ensues [254]. The capacity of extremely thermoacidophilic archaea, involved in biomining, to oxidize arsenite differentiates them from the majority of their mesophilic counterparts [255].Many phylogenetically distinct bacteria are known to oxidize As3+ [256,257] using the heterodimeric enzyme Aio (formerly Aox, Aro or Aso; see [258]), comprised of the AioA (molybdopterin) and AioB (Rieske) subunits. Among archaea with sequenced genomes, which does not include S. metallicus or A. brierleyi, several including Pyrobaculum calidifontis, Sulfolobus tokodaii, and Aeropyrum pernix, were found to harbor aio clusters, indicating these as putative As3+-oxidizing archaea [256]. The aio gene cluster appears to be linked to an “ancient” bioenergetic pathway [257].
In general, few studies exist exploring cadmium resistance mechanisms among acidophiles, especially extreme thermoacidophiles. The common detoxification mechanism in neutrophiles employs a variety of active efflux systems [144,145]. Although not elucidated, analysis of sequenced acidophile genomes indicates cadmium efflux, mediated by CadA, might be a common resistance mechanism amongst acidophiles [259]. Exposure of S. metallicus to cadmium revealed the response of two cop loci, suggesting the locus not only functions for copper detoxification, but cadmium as well [204]. Additionally, the cadmium response, along with copper, elicited a defensive stress response including proteins related production and conversion of energy, amino acids biosynthesis, stress responses, and transcription regulation. The results of a general defensive response are consistent with previous characterization and appear to represent a general cellular response to metal challenge [203,242,260,261].
No determinants of nickel resistance have been experimentally identified in acidophilic archaea, despite a detoxification system, based on efflux, existing for bacteria [144,145]. The nickel resistance determinant has been identified for acidophilic bacteria Leptospirillum ferriphilum, which was attributed to a nickel–cobalt resistance operon (NCR) [262,263]. However, the only study to date in acidophilic archaea identified redox stress proteins involved in the adaptation response of S. solfataricus to nickel challenge [260].
A number of processes have been investigated for the bioaccumulation of uranium, which includes biosorption [264], bioreduction [265], and biominerlization [266,267,268]. These studies have largely focused on bacteria with the mechanisms of uranium accumulation and the resulting uranium complexes being poorly understood in archaea. Given the differences in cell wall structures between archaea and bacteria, differences in interaction mechanisms can be expected [269]. The anaerobic hyperthermophile, Pyrobaculum islandicum, has been shown to reduce U(VI) to the insoluble U(IV) mineral uraninite leading to the formation of extracellular deposits [270]. Dense uranium deposits were observed at the cell surface in the halophilic archaeon Halobacterium halobium, with complexation of uranium predominantly via cellular inorganic phosphate (uranyl phosphate) [271]. More recently, the interaction of S. acidocaldarius with U(VI) was studied under highly acidic (pH 1.5–3.0) and moderately acidic (pH 4.5) conditions, relevant to the physiological growth optimum of this organism and uranium polluted environments [269,272]. For the highly acidic conditions, U(VI) was demonstrated to complex with organic phosphate groups, while under moderately acid conditions carboxylic groups were also involved in U(VI) complexation. Intracellular deposits associated with the inner side of the cytoplasmic membrane represented the majority of U(VI) accumulation, with a small amount biomineralized extracellularly [269]. In contrast to the use of organic phosphate groups, neutrophilic bacteria are known to secrete orthophosphate (via polyphosphate metabolism) and form inorganic uranium precipitates that serve to protect bacterial cells from uranium toxicity [266,267,273]. The pH dependence of uranium complexation in S. acidocaldarius differs from the pH independent process of the acidophilic bacterium A. ferrooxidans, where uranium complexation occurred solely via organic phosphate groups between pH 2–4.5 [274,275]. Further, in contrast to U(VI) biosorption in Chryseomonas sp. [276], Bacillus sphaericus ATCC 14577 [277], Pseudomonas fluorescence ATCC 55241 [271], and H. halobium [271] under corresponding experimental conditions, S. acidocaldarius has a significantly lower capability [269]. As noted above, bacteria have significantly different cell wall structures from archaea, which contain a large number of uranium binding ligands, such as carboxylic and phosphate groups [169,278]. Although the cell wall of H. halobium only consists of a S-layer protein, in contrast to S. acidocaldarius this S-layer is enriched in carboxylic amino acid residues and can explain the higher uranium binding capacity [269,279]. A few microorganisms, such as like Bacillus sphaericus JG-A12, possess a phosphorylated S-layer allowing for binding of large amounts of uranium [169]. Taken together, S. acidocaldarius appears to interact with and detoxify U(VI) differently than other acidophilic and non-acidophilic bacteria. Recently, a preliminary investigation into the uranium resistance of M. prunae compared to M. sedula indicated a novel role of a toxin-antitoxin in resistance [36].

5. Attachment of Extreme Thermoacidophiles to Surfaces

The vast majority of leaching bacteria adhere to the mineral sulfide surface, generally mediated by an exopolysaccharide (EPS) surrounding the cells [280,281,282,283,284,285]. The EPS provides an essential micro-environment and reaction space for organisms leaching mineral sulfides [286,287,288,289]. Certain species, like Acidithiobacillus caldus, cannot adhere and requires co-culture with EPS-forming acidophiles [290]. Curiously, if an organism is capable of mineral sulfide attachment, the space for attachment must be non-limiting [291,292]. The formation of EPS is known to be stimulated by attachment or surface contact [285,293]. The composition of EPS consists of sugars, fatty acids, glucuronic acid, and Fe(III) ions [39,281,294]. Adherence is mainly attributed to electrostatic interactions, but hydrophobic interactions do contribute and the magnitude of the adhesion force has been determined [281,295,296,297,298,299]. The EPS does display adaptability depending on whether the substrate is a metal sulfide or sulfur [281], though the molecular mechanisms used to adapt composition and amount of EPS according to growth substrate are still unknown [291]. While the site of attachment and mechanisms for specific site detection are still unknown, the process does not appear to be random, with cells attaching to areas of surface imperfection or low-degree of crystallinity [281,291,294,300,301,302,303,304]. Additional elements mediating adherence to surfaces include pili and S-layer proteins [170,305,306,307].
L. ferrooxidans and A. ferrooxidans possess chemotaxis systems for sensing Fe(II), which might function to identify specific sites on pyrite surfaces for attachment [308,309]. Quorum sensing functions in bioleaching bacterium allow for swarming behavior on metal sulfides and play a key role in biofilm formation, enhancing dissolution of the mineral substrate [310,311,312,313,314,315]. Early biofilm formation involves capsular polysaccharide production (CPS), up-regulation of genes related to pili and EPS production, motility and quorum sensing, synthesis of cell wall structures, specific proteases, stress response chaperons, and mixed acid fermentation [316,317,318,319]. Proteomic analysis revealed similar results with the addition of increased production of osmolarity sensing, outer-membrane efflux, iron uptake, sulfate uptake and assimilation, glutathione/coenzyme/cofactor biosynthesis, lipoproteins, and nucleotidases [320].
As discussed above, chalcopyrite bioleaching is more effective at temperatures above 65 °C, requiring the use of extremely thermoacidophilic microorganisms [4,321,322,323]. The majority of studies, related to mineral sulfides, focused on attachment parameters of temperature and culture history, the influence of planktonic and attached cells on the dissolution process, visualizing pyrite leaching, and biofilm development [324,325,326,327]. As is the case for other acidophilic metal mobilizers, EPS plays an important role in the adhesion to solid mineral substrates for extreme thermoacidophiles [34,39,283,328]. The EPS for S. acidocaldarius, S. solfataricus, and S. tokodaii, contains mannose, galactose, and N-acetylglucosamine [328,329]. Thicknesses of EPS for M. hakonensis grown on pyrite and chalcopyrite have been determined to be 8–12 and 4–8 μm, respectively [283]. Elemental analysis of EPS produced by M. hakonensis, grown on chalcopyrite, showed iron levels below the detection limit, preventing assessment of the presence or absence of iron in the EPS [39]. The result suggests differences in Fe(II)-oxidation enzymes could be more important for dissolution than iron levels in the EPS. For S. solfataricus and S. acidocaldarius and likely other extremely thermoacidophilic archaea, initial attachment to solid substrates involves pili, and additionally flagella for S. solfataricus [328,330,331].
A proteomic/transcriptomic study of S. acidocaldarius, S. solfataricus, and S. tokodaii adjustment to biofilm lifestyle was strain specific [329]. As noted above for other acidophiles, these changes were largely associated with energy production and conversion, amino acid metabolism, lipid and carbohydrate metabolism, transport related functions, and cell surface modifications. Interestingly, very few changes were shared across the species, which included a family of Lrs14-like transcriptional regulators, several significantly influencing biofilm formation or cell motility [332]. No quorum sensing (QS)-phenomena were detected in the biofilm formation of S. acidocaldarius, S. solfataricus, and S. tokodaii nor for F. acidarmanus Fer1, leaving the significance of cell signaling and communication unknown [329,333]. The biofilms formed by F. acidarmanus rely on EPS and involve shifts in metabolism towards anaerobic growth. Further, the biofilms are monolayer, and like acidophilic bacteria, appear to preferentially occur at cracks/defects on pyrite surfaces [284,333]. Recently, a methodology for investigating archaeal biofilms was developed using fluorescence lectin-binding analysis. Results showed variations in EPS glycoconjugates for three archaeal species and that various substrates induce different EPS glycoconjugates, similar to the flexibility of bacteria [281,334]. For more information on the aspects of other archaeal biofilms, informative reviews are available [331,335,336,337,338].
In general, there exist three models describing microbe-mineral electron transfer: (i) direct; (ii) electron shuttle; (iii) nanowire [339,340]. For metal-reducing Shewanella and Geobacter species, several strategies have been proposed to mediate interfacial electron transport from the cell to the external solid-phase electron sink, though intense debate still exits concerning molecular details. For short distances, <2 nm, electron tunneling could play a critical role in electron transfer, whereby direct electron transfer occurs between the extracellular substrate and redox-active enzyme [341,342,343]. However, dramatically longer distances of electron transfer have been reported, ranging from nanometers to centimeters, requiring long-distance electron transport models [344]. A model for diffusive shuttling of electrons involves flavin-mediated transfer of electrons between the extracellular substrate and redox-active enzymes, multi-haem cytochromes, on the cell surface [345,346,347]. A model based on extracellular appendages, commonly called nanowires, involves electron transfer along these nanowires, believed to be either membrane- or pilin-based, between the solid substrate and cell [265,348,349,350,351,352]. Interestingly, the occurrence of nanowires coincides with formation of separate or attached redox-active membrane vesicles [352,353]. Further, bacterial biofilms incorporating nanowires or outer membrane cytochromes and multicellular bacterial cables can transfer electrons over long distances [354,355]. Electron conductance is proposed to occur from either metallic-like band transport or multi-step redox hopping mechanisms, though the former remains controversial [356,357,358,359,360].
A tentative “contact” model for metal sulfide dissolution (e.g., FeS2), requiring an oxidizing attack by Fe(III), has been proposed [291]. The model postulates that Fe(III), complexed to glucuronic acid in the EPS, performs the oxidizing attack of the metal sulfide. The Fe(II) produced by the cathodic electron transfer is then released from EPS chelators and diffuses toward the outer membrane where (re)oxidation occurs, thus the cycle repeats [291]. The model is similar to the flavin-mediated shuttling, noted above, with Fe(II) serving as the electron shuttle. The metal sulfide model rests on four assumptions: (i) oxidizing attack by Fe(III) is required; (ii) EPS-complexed Fe(III) fulfills this function; (iii) electron tunneling effects explain transfer, and (iv) Fe(II) ion-glucuronic acid complexes are less stable than Fe(III). The first assumption rests on the assertion that direct electron transfer from the metal sulfide to the attached cell does not occur, since no enzymes or nanowires have been demonstrated for metal sulfide attached cells [291]. This assumption seems tenuous, given the recent work on metal reducing species showing usage of nanowires for metal reduction, discussed above. Additionally, type IV pili in A. ferrooxidans are highly conductive and might function as nanowires, directly transferring electrons from the external substrate [361]. The second assumption is based on experimental evidence where an A. ferrooxidans strain with high Fe(III) in the EPS had a higher bioleaching capacity compared to strains with low Fe(III) concentration [289,294]. A similar phenomenon has been seen for L. ferrooxidans, but the results only revealed the importance of the local concentration of the corrosion promotor Fe(III) in the biofilm environment [362]. The complexation “probably” occurs with glucuronic acid residues, but conclusive evidence along with Fe(II)/Fe(III) binding constants has not been presented [288,289,294]. Electron tunneling over distances <2 nm is widely accepted [341,363] and would allow both EPS complexed and solution Fe(III) to be reduced. However, given the thickness of the EPS is 10–100 nm wide, not all Fe(III) would be within range, requiring diffusion towards the surface [286,364].
The above model does not incorporate the potential for mineral sulfide destabilizers that could help initiate release of Fe(II). Cysteine is known to accelerate FeS2 dissolution, possibly by disrupting the FeS2 surface, causing release of iron-sulfur species [365,366]. Further, A. ferrooxidans’ aporusticyanin was suspected to function as a receptor for initial adhesion to mineral sulfides, in which the protein could destabilize the mineral surface, leading to Fe(II) dissolution [367].
As mentioned above, the occurrence of nanowires coincides with formation of separate or attached redox-active membrane vesicles for the metal-reducing Shewanella species [352,353]. In bacteria, vesicles have roles in colonization and cell co-aggregation, both critical to biofilm formation [368]. Gram-negative bacteria, along with the extremely thermoacidophilic archaea S. acidocaldarius, S. solfataricus, S. tokodaii and S. islandicus, release membrane vesicles [369,370,371,372]. The presence of archaeal homologous of the eukaryotic endosomal sorting complex required for transport-I (ESCRT) proteins in the crenarchaeal vesicles, suggests vesicle formation occurs through an outward budding process, similar to inward budding of the endosomal compartment in eukaryotes [372,373]. The archaeal vesicles could serve a homologous function in electron transfer from the solid substrate to the cell, as for the metal reducers.
The possibility that multiple mechanisms of interaction occur throughout different stages of mineral oxidation seems possible [374]. Initially, cells localize to non-random sites on the mineral sulfide surface, through an unknown mechanism, and attachment proceeds by CPS, pili, flagella, S-layer, mineral receptors (e.g., aporusticyanin), or a combination. Once attached mineral destabilizers cause an initial release of iron-sulfur species and cells switch to a sessile growth mode. EPS production occurs, providing an essential micro-environment and reaction space, where the corrosion promoter Fe(III) is entrapped and accelerates mineral dissolution. Though not experimentally observed to date, the implications of conductive pili and redox-active vesicles should not be ruled out.

6. Bioleaching

Over the course of the past few decades, biomining has centered on the development of technologies to recover precious metals contained within ore-bearing matrices. In the recent past, numerous industrial processes have matured, primarily those involving the recovery of gold from refractory ores or the recovery of nickel or copper from base metal sulfides. In fact, some estimates suggest that as much as 15% of copper and 5% of gold production (on a global scale) utilize microbial-assisted extraction technologies [375]. Further, as the relative availability of higher-grade ores diminishes and environmental regulation increases, it is likely that interest in biomining will increase in an attempt to improve metal selectivity and yield, while minimizing the release of toxic pollutants.
Recent reviews have emphasized the importance of mesophilic and moderately thermophilic acidophiles involved in bioleaching, ranging from industrial prospects [292,323,375] to specific uses for secondary copper ores [376] and polymetallic ores [377]. There are fewer details on the successful development of bioleaching applications using extremely thermoacidophilic microorganisms [378]. Extremely thermoacidophilic bioleaching, as it exists, is dominated by the genera Acidianus, Metallosphaera, and Sulfolobus for copper recovery from recalcitrant ores [379,380] or for sulfur oxidation to improve gold recovery from biooxidation, e.g., BIOPROTM [381].

6.1. Current Biooxidation/Bioleaching Practices at Elevated Temperatures

The treatment of primary copper ores, such as chalcopyrite (CuFeS2), has been the main driver of extremely thermoacidophilic bioleaching developments, see Figure 4. Under mesophilic conditions, heap bioleaching of chalcopyrite tends to achieve low copper yields, often attributed to passivation, or the formation of deposited layers of iron complexes or polysulfides on the mineral surface [323,375,376]. These passivation effects do not appear to be as severe in extremely thermoacidophilic cultures, based on laboratory evaluations and pilot plant testing demonstrating high copper dissolution [9,379,380,382]. Although kinetics may be a driving factor in the dissolution process, evidence has emerged that redox potential can play a role in mitigating passivation for some circumstances [383,384]. In fact, it may be possible to greatly improve the metal dissolution of mesophilic and moderately thermophilic organisms by redox controlling strategies [384] or by the addition of silver, which forms a galvanic couple in the presence of chalcopyrite [385]. However, this result may support a more recent hypothesis that accounts for lowered dissolution of chalcopyrite, due to electronic and interfacial structure that more closely resembles a semiconductor [386]. Given that iron precipitates and polysulfides are so commonplace in both successful and unsuccessful leaching operations, more research is needed to understand if the intrinsic difference in dissolution rates is related to temperature/kinetics or perhaps a yet to be discovered dissolution mechanism among the extreme thermoacidophiles.
Extremely thermoacidophilic archaea have some niche advantages in copper biomining. In the case of chalcopyrite-bearing molybdenite, extreme thermoacidophiles achieve selective copper dissolution, leading to improvement of molybdenum flotation concentrates, with minimal (<10%) molybdenum dissolution [387,388]. Further, there is evidence that extremely thermoacidophilic archaea may be more adept than mesophilic bacteria at extracting copper from other copper-sulfide ores, based on column bioleaching involving covellite (CuS) and enargite (Cu3AsS4) [6]. This suggests their potential applications in processing mixed copper-bearing ores at elevated temperatures. Also, a study involving A. brierley bioleaching of enargite showed that the species can selectively mobilize copper, while simultaneously precipitating arsenic in the form of arsenate [389]. This approach could limit the deleterious effects of mining ores containing arsenic.
Figure 4. Chronology of selected developments in extreme thermoacidophilic biotechnology [6,9,32,53,54,379,380,382,384,390,391,392,393,394,395,396,397,398,399,400].
Figure 4. Chronology of selected developments in extreme thermoacidophilic biotechnology [6,9,32,53,54,379,380,382,384,390,391,392,393,394,395,396,397,398,399,400].
Minerals 05 00397 g004
Issues with sulfur oxidation have emerged in some biooxidation/bioleaching processes. In particular, gold and copper biomining of ores containing high concentrations of pyrite/pyrrhotite presents a challenge in mesophilic bioleaching processes. In the case of gold biomining, sulfur is often retained in a partially oxidized form (such as elemental sulfur). This residual sulfur is then capable of reacting with cyanide in downstream processing, severely impacting recoveries and increasing operating costs [381,401]. A consortium of mesophilic, moderately thermophilic, and extremely thermoacidophilic microbes has shown promise in improving sulfur oxidation and subsequent metal recoveries [381,401]. In the case of pyrite/pyrrhotite-rich copper deposits, heap bioleaching generates large amounts of heat [401,402]. In some instances, the inability to control temperature in the heap, due to the exothermic nature of sulfur oxidation, can result in issues with population succession and dissolution [401,402]. This issue might be mitigated by the use of extremely thermoacidophilic archaea. However, column leaching experiments involving copper ores have revealed a tendency to form percolation channels [6,9]. This is possibly due to iron precipitation as oxyhydroxysulfates at higher temperatures [10]. To improve the efficacy of extremely thermoacidophilic organisms bioleaching copper in heaps, one technology, GEOCOAT®, utilizes ground copper-ore concentrate coated onto a barren rock surface to increase surface area and, as a consequence, achieve increased rates and overall dissolution of copper [397,403].

6.2. Extreme Thermoacidophile Process Challenges

Currently, extremely thermoacidophilic bioleaching presents certain process dynamic challenges. One potential concern is the delicate nature of the archaeal cell envelope, which lacks the bacterial peptidoglycan outer-membrane. This potentially places a limit on agitation rates that the microbes can endure in tank bioleaching conditions and may facilitate the need for highly specialized turbine/agitator designs [378,379,380]. Another issue is oxygen demand in high temperature environments. A well-known consequence of higher temperatures is decreased dissolved oxygen content, requiring the use of enriched oxygen sources (at a much higher operating costs than air) [378,379,380]. Compounding this issue is the production of reactive oxygen species, especially in the presence of finely ground mineral stocks and low pH, conditions which typically optimize leaching [404,405]. In fact, this result may be a critical issue that prevents higher solids loading in extremely thermoacidophilic bioreactors [405]. However, in all of these cases, the concerns raised for extreme thermoacidophiles as bioleachers have not been confirmed, but should be assessed as related technologies move forward.

6.3. Polymetallic Ores and Industrial Waste

A common issue in modern mining is the need to maximize yields of numerous metals of varying values from complex polymetallic ores. Several studies over the past few decades have suggested the potential value of bioleaching complex deposits with extreme thermoacidophiles. In the case of zinc, high recovery has been observed from complex sulfides, containing sphalerite [54,396]. In this instance, the extreme thermophiles appear to outperform moderate thermophiles and mesophiles. More recent studies continue to highlight the ability of extreme thermoacidophiles to leach a variety of metals from complex ores. In the case of a black shale (containing Mn, Fe, Zn, Ni, Cu, and Co), over 90% dissolution of manganese, copper, zinc, and nickel was achieved with extreme thermoacidophilic cultures [406]. In the case of ores containing chalcopyrite, sphalerite, and galena, greater than 90% dissolution of copper and zinc were observed in the leachate, with more than 90% recovery of lead following brine precipitation [400]. Thus, the potential for improved kinetics of bioleaching by extreme thermoacidophiles is not limited to the current narrowly applied fields of sulfur oxidation and copper dissolution.
Another area of growing interest is the treatment of industrial and consumer waste [407,408]. A. brierleyi has been used to treat spent hydrocatalysts from petroleum processing in order to remove the molybdenum and nickel from the catalyst prior to disposal [399]. Another study focused on remediation of mining spillage composed primarily of pyrite, with some sphalerite and arsenopyrite. In this instance, extreme thermoacidophiles showed much faster kinetics for the dissolution of iron, zinc, and arsenic [398].

7. Conclusions

In general, bioleaching is likely to remain an important avenue for recovery of base, precious and strategic metals from mining operations. This processing approach reflects a trend toward more stringent environmental regulations which are incentivizing the use of sustainable industrial practices. In addition, the depletion of high-grade ores and the need to process increasing amounts of heavy metal waste will inevitably create a processing bottleneck, if only conventional chemical/physical metal extraction techniques are considered. Looking further into the future, implications for using extremely thermoacidophiles in asteroid mining creates yet another technological dimension [409].
Given the potential advantages and challenges associated with high temperature bioleaching operations, efforts to further understand the underlying metabolic, physiological and genetic mechanisms characteristic of extreme thermoacidophiles need to continue. In particular, as molecular genetics tools become more tractable and allow for metabolic engineering of biomining microorganisms to improve their efficacy, the corresponding issues with release of genetically modified organisms (GMOs) also arise. However, the unique aspects of extreme thermoacidophiles, and the inhospitable nature of biomining sites, may mitigate some of the concerns normally associated with release of GMOs. These concerns will need to be addressed from the perspective of microbial ecology of hot acid biotopes.
A new and exciting frontier, of strategic importance where bioleaching microorganisms could play a significant role, is in the extraction and recovery of rare earth elements. Microorganisms and certain fungi can accumulate and absorb rare earth elements, providing a fundamental framework to build novel extraction and recovery processes [410,411,412,413]. Interestingly, an extremely acidophilic methanotrophic microorganism requires certain rare earth elements for survival [414]. Given that microbial metalloproteomes are largely uncharacterized, the potential for discovering novel rare earth element binding proteins among extreme thermoacidophiles is promising [415].


This work was supported in part by the U.S. Defense Threat Reduction Agency (DTRA) (Grant No. HDTRA1-09-0030) and the U.S. Air Force Office of Scientific Research (AFOSR) (FA9550-13-1-0236).

Author Contributions

Garrett Wheaton, James Counts, Arpan Mukherjee, Jessica Kruh and Robert Kelly contributed to the writing and analysis of the material included in this review.

Conflicts of Interest

The authors declare no conflict of interest.


  1. Brierley, J.A.; Brierley, C.L. Present and future commercial applications of biohydrometallurgy. Hydrometallurgy 2001, 59, 233–239. [Google Scholar] [CrossRef]
  2. Rawlings, D.E.; Dew, D.; du Plessis, C. Biomineralization of metal-containing ores and concentrates. Trends Biotechnol. 2003, 21, 38–44. [Google Scholar] [CrossRef]
  3. Bhakta, P.; Arthur, B. Heap bio-oxidation and gold recovery at newmont mining: First-year results. J. Miner. Met. Mater. Soc. 2002, 54, 31–34. [Google Scholar] [CrossRef]
  4. Rawlings, D.E. Heavy metal mining using microbes. Annu. Rev. Microbiol. 2002, 56, 65–91. [Google Scholar] [CrossRef] [PubMed]
  5. Rawlings, D.E. Characteristics and adaptability of iron- and sulfur-oxidizing microorganisms used for the recovery of metals from minerals and their concentrates. Microb. Cell Fact. 2005, 4. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  6. Lee, J.; Acar, S.; Doerr, D.L.; Brierley, J.A. Comparative bioleaching and mineralogy of composited sulfide ores containing enargite, covellite and chalcocite by mesophilic and thermophilic microorganisms. Hydrometallurgy 2011, 105, 213–221. [Google Scholar] [CrossRef]
  7. Norris, P.R.; Burton, N.P.; Foulis, N.A. Acidophiles in bioreactor mineral processing. Extremophiles 2000, 4, 71–76. [Google Scholar] [CrossRef] [PubMed]
  8. Acar, S.; Brierley, J.A.; Wan, R.Y. Conditions for bioleaching a covellite-bearing ore. Hydrometallurgy 2005, 77, 239–246. [Google Scholar] [CrossRef]
  9. Norris, P.R.; Calvo-Bado, L.A.; Brown, C.F.; Davis-Belmar, C.S. Ore column leaching with thermophiles: I, copper sulfide ore. Hydrometallurgy 2012, 127, 62–69. [Google Scholar] [CrossRef]
  10. Norris, P.R.; Brown, C.F.; Caldwell, P.E. Ore column leaching with thermophiles: II, polymetallic sulfide ore. Hydrometallurgy 2012, 127, 70–76. [Google Scholar] [CrossRef]
  11. Norris, P.R.; Burton, N.P.; Clark, D.A. Mineral sulfide concentrate leaching in high temperature bioreactors. Miner. Eng. 2013, 48, 10–19. [Google Scholar] [CrossRef]
  12. Pradhan, N.; Nathsarma, K.C.; Srinivasa Rao, K.; Sukla, L.B.; Mishra, B.K. Heap bioleaching of chalcopyrite: A review. Miner. Eng. 2008, 21, 355–365. [Google Scholar] [CrossRef]
  13. Schippers, A. Microorganisms involved in bioleaching and nucleic acid-based molecular methods for their identification and quantification. In Microbial Processing of Metal Sulfides; Donati, E.R., Sand, W., Eds.; Springer: Dordrecht, The Netherlands, 2007; pp. 3–33. [Google Scholar]
  14. Norris, P.R. Acidophile Diversity in Mineral Sulfide Oxidation. In Biomining; Rawlings, D., Johnson, D.B., Eds.; Springer: Berlin, Germany, 2007; pp. 199–216. [Google Scholar]
  15. Ilbert, M.; Bonnefoy, V. Insight into the evolution of the iron oxidation pathways. Biochim. Biophys. Acta 2013, 1827, 161–175. [Google Scholar] [CrossRef] [PubMed]
  16. Brock, T.D.; Brock, K.M.; Belly, R.T.; Weiss, R.L. Sulfolobus: A new genus of sulfur-oxidizing bacteria living at low pH and high temperature. Arch. Mikrobiol. 1972, 84, 54–68. [Google Scholar] [CrossRef] [PubMed]
  17. Grogan, D.W. Phenotypic characterization of the archaebacterial genus Sulfolobus: Comparison of five wild-type strains. J. Bacteriol. 1989, 171, 6710–6719. [Google Scholar] [PubMed]
  18. Huber, G.; Drobner, E.; Huber, H.; Stetter, K.O. Growth by Aerobic Oxidation of Molecular Hydrogen in Archaea—A Metabolic Property so far Unknown for this Domain. Syst. Appl. Microbiol. 1992, 15, 502–504. [Google Scholar] [CrossRef]
  19. Takayanagi, S.; Kawasaki, H.; Sugimori, K.; Yamada, T.; Sugai, A.; Ito, T.; Yamasato, K.; Shioda, M. Sulfolobus hakonensis sp. nov., a novel species of acidothermophilic archaeon. Int. J. Syst. Bacteriol. 1996, 46, 377–382. [Google Scholar] [CrossRef] [PubMed]
  20. Chen, L.; Brügger, K.; Skovgaard, M.; Redder, P.; She, Q.; Torarinsson, E.; Greve, B.; Awayez, M.; Zibat, A.; Klenk, H.; et al. The genome of Sulfolobus acidocaldarius, a model organism of the Crenarchaeota. J. Bacteriol. 2005, 187, 4992–4999. [Google Scholar] [CrossRef] [PubMed]
  21. Zillig, W.; Stetter, K.O.; Wunderl, S.; Schulz, W.; Priess, H.; Scholz, I. The Sulfolobus-“Caldariella” group: Taxonomy on the basis of the structure of DNA-dependent RNA polymerases. Arch. Microbiol. 1980, 125, 259–269. [Google Scholar] [CrossRef]
  22. She, Q.; Singh, R.K.; Confalonieri, F.; Zivanovic, Y.; Allard, G.; Awayez, M.J.; Chan-Weiher, C.C.; Clausen, I.G.; Curtis, B.A.; de Moors, A.; et al. The complete genome of the crenarchaeon Sulfolobus solfataricus P2. Proc. Natl. Acad. Sci. USA 2001, 98, 7835–7840. [Google Scholar] [CrossRef] [PubMed]
  23. Grogan, D.; Palm, P.; Zillig, W. Isolate B12, which harbours a virus-like element, represents a new species of the archaebacterial genus Sulfolobus, Sulfolobus shibatae, sp. nov. Arch. Microbiol. 1990, 154, 594–599. [Google Scholar] [CrossRef] [PubMed]
  24. Huber, G.; Stetter, K.O. Sulfolobus metallicus, sp. nov., a novel strictly chemolithoautotrophic thermophilic archaeal species of metal-mobilizers. Syst. Appl. Microbiol. 1991, 14, 372–378. [Google Scholar] [CrossRef]
  25. Bathe, S.; Norris, P.R. Ferrous iron- and sulfur-induced genes in Sulfolobus metallicus. Appl. Environ. Microbiol. 2007, 73, 2491–2497. [Google Scholar] [CrossRef] [PubMed]
  26. Suzuki, T.; Iwasaki, T.; Uzawa, T.; Hara, K.; Nemoto, N.; Kon, T.; Ueki, T.; Yamagishi, A.; Oshima, T. Sulfolobus tokodaii sp. nov. (f. Sulfolobus sp. strain 7), a new member of the genus Sulfolobus isolated from Beppu Hot Springs, Japan. Extremophiles 2002, 6, 39–44. [Google Scholar]
  27. Inatomi, K.; Ohba, M.; Oshima, T. Chemical properties of proteinaceus cell wall from an acido-thermophile, Sulfolobus acidocaldarius. Chem. Lett. 1983, 12, 1191–1194. [Google Scholar] [CrossRef]
  28. Kawarabayasi, Y.; Hino, Y.; Horikawa, H.; Jin-no, K.; Takahashi, M.; Sekine, M.; Baba, S.; Ankai, A.; Kosugi, H.; Hosoyama, A.; et al. Complete genome sequence of an aerobic thermoacidophilic crenarchaeon, Sulfolobus tokodaii strain7. DNA Res. 2001, 8, 123–140. [Google Scholar] [CrossRef] [PubMed]
  29. Jan, R.L.; Wu, J.; Chaw, S.M.; Tsai, C.W.; Tsen, S.D. A novel species of thermoacidophilic archaeon, Sulfolobus yangmingensis sp. nov. Int. J. Syst. Bacteriol. 1999, 49, 1809–1816. [Google Scholar] [CrossRef] [PubMed]
  30. Xiang, X.; Dong, X.; Huang, L. Sulfolobus tengchongensis sp. nov., a novel thermoacidophilic archaeon isolated from a hot spring in Tengchong, China. Extremophiles 2003, 7, 493–498. [Google Scholar] [CrossRef] [PubMed]
  31. Zillig, W.; Kletzin, A.; Schleper, C.; Holz, I.; Janekovic, D.; Hain, J.; Lanzendörfer, M.; Kristjansson, J.K. Screening for Sulfolobales, their Plasmids and their Viruses in Icelandic Solfataras. Syst. Appl. Microbiol. 1994, 16, 609–628. [Google Scholar] [CrossRef]
  32. Huber, G.; Spinnler, C.; Gambacorta, A.; Stetter, K. Metallosphaera sedula gen, and sp. nov. represents a new genus of aerobic, metal-mobilizing, thermoacidophilic archaebacteria. Syst. Appl. Microbiol. 1989, 12, 38–47. [Google Scholar] [CrossRef]
  33. Auernik, K.S.; Kelly, R.M. Identification of components of electron transport chains in the extremely thermoacidophilic crenarchaeon Metallosphaera sedula through iron and sulfur compound oxidation transcriptomes. Appl. Environ. Microbiol. 2008, 74, 7723–7732. [Google Scholar] [CrossRef] [PubMed]
  34. Auernik, K.S.; Maezato, Y.; Blum, P.H.; Kelly, R.M. The genome sequence of the metal-mobilizing, extremely thermoacidophilic archaeon Metallosphaera sedula provides insights into bioleaching-associated metabolism. Appl. Environ. Microbiol. 2008, 74, 682–692. [Google Scholar] [CrossRef] [PubMed]
  35. Fuchs, T.; Huber, H.; Teiner, K.; Burggraf, S.; Stetter, K.O. Metallosphaera prunae, sp. nov., a novel metal-mobilizing, thermoacidophilic archaeum, isolated from a uranium mine in Germany. Syst. Appl. Microbiol. 1995, 18, 560–566. [Google Scholar] [CrossRef]
  36. Mukherjee, A.; Wheaton, G.H.; Blum, P.H.; Kelly, R.M. Uranium extremophily is an adaptive, rather than intrinsic, feature for extremely thermoacidophilic Metallosphaera species. Proc. Natl. Acad. Sci. USA 2012, 109, 16702–16707. [Google Scholar] [CrossRef] [PubMed]
  37. Kurosawa, N. Reclassification of Sulfolobus hakonensis Takayanagi et al. 1996 as Metallosphaera hakonensis comb. nov. based on phylogenetic evidence and DNA G+C content. Int. J. Syst. Evol. Microbiol. 2003, 53, 1607–1608. [Google Scholar] [CrossRef] [PubMed]
  38. Plumb, J.J.; Muddle, R.; Franzmann, P.D. Effect of pH on rates of iron and sulfur oxidation by bioleaching organisms. Miner. Eng. 2008, 21, 76–82. [Google Scholar] [CrossRef]
  39. Usher, K.M.; Shaw, J.A.; Kaksonen, A.H.; Saunders, M. Elemental analysis of extracellular polymeric substances and granules in chalcopyrite bioleaching microbes. Hydrometallurgy 2010, 104, 376–381. [Google Scholar] [CrossRef]
  40. Liu, L.-J.; You, X.-Y.; Guo, X.; Liu, S.-J.; Jiang, C.-Y. Metallosphaera cuprina sp. nov., an acidothermophilic, metal-mobilizing archaeon. Int. J. Syst. Evol. Microbiol. 2011, 61, 2395–2400. [Google Scholar] [CrossRef] [PubMed]
  41. Liu, L.-J.; You, X.-Y.; Zheng, H.; Wang, S.; Jiang, C.-Y.; Liu, S.-J. Complete genome sequence of Metallosphaera cuprina, a metal sulfide-oxidizing archaeon from a hot spring. J. Bacteriol. 2011, 193, 3387–3388. [Google Scholar] [CrossRef] [PubMed]
  42. Kozubal, M.; Macur, R.E.; Korf, S.; Taylor, W.P.; Ackerman, G.G.; Nagy, A.; Inskeep, W.P. Isolation and distribution of a novel iron-oxidizing crenarchaeon from acidic geothermal springs in Yellowstone National Park. Appl. Environ. Microbiol. 2008, 74, 942–949. [Google Scholar] [CrossRef] [PubMed]
  43. Kozubal, M.A.; Dlakic, M.; Macur, R.E.; Inskeep, W.P. Terminal oxidase diversity and function in “Metallosphaera yellowstonensis”: Gene expression and protein modeling suggest mechanisms of Fe(II) oxidation in the sulfolobales. Appl. Environ. Microbiol. 2011, 77, 1844–1853. [Google Scholar] [CrossRef] [PubMed]
  44. Bettstetter, M.; Peng, X.; Garrett, R.A.; Prangishvili, D. AFV1, a novel virus infecting hyperthermophilic archaea of the genus acidianus. Virology 2003, 315, 68–79. [Google Scholar] [CrossRef]
  45. Basta, T.; Smyth, J.; Forterre, P.; Prangishvili, D.; Peng, X. Novel archaeal plasmid pAH1 and its interactions with the lipothrixvirus AFV1. Mol. Microbiol. 2009, 71, 23–34. [Google Scholar] [CrossRef] [PubMed]
  46. You, X.Y.; Liu, C.; Wang, S.Y.; Jiang, C.Y.; Shah, S.A.; Prangishvili, D.; She, Q.; Liu, S.J.; Garrett, R.A. Genomic analysis of Acidianus hospitalis W1 a host for studying crenarchaeal virus and plasmid life cycles. Extremophiles 2011, 15, 487–497. [Google Scholar] [CrossRef] [PubMed]
  47. Giaveno, M.A.; Urbieta, M.S.; Ulloa, J.R.; González Toril, E.; Donati, E.R. Physiologic Versatility and Growth Flexibility as the Main Characteristics of a Novel Thermoacidophilic Acidianus Strain Isolated from Copahue Geothermal Area in Argentina. Microb. Ecol. 2013, 65, 336–346. [Google Scholar] [CrossRef] [PubMed]
  48. Urbieta, M.S.; Rascovan, N.; Castro, C.; Revale, S.; Giaveno, M.A.; Vazquez, M.; Donati, R. Draft Genome Sequence of the Novel Thermoacidophilic Archaeon Acidianus copahuensis Strain ALE1, Isolated from the Copahue Volcanic Area in Neuquén, Argentina. Genome Announc. 2014, 2. [Google Scholar] [CrossRef] [PubMed]
  49. Segerer, A.; Neuner, A.; Kristjansson, J.K.; Stetter, K.O. Acidianus infernus gen. nov., sp. nov., and Acidianus brierleyi Comb. nov.: Facultatively aerobic, extremely acidophilic thermophilic sulfur-metabolizing archaebacteria. Int. J. Syst. Bacteriol. 1986, 36, 559–564. [Google Scholar] [CrossRef]
  50. Plumb, J.J.; Haddad, C.M.; Gibson, J.A.E.; Franzmann, P.D. Acidianus sulfidivorans sp. nov., an extremely acidophilic, thermophilic archaeon isolated from a solfatara on Lihir Island, Papua New Guinea, and emendation of the genus description. Int. J. Syst. Evol. Microbiol. 2007, 57, 1418–1423. [Google Scholar] [CrossRef] [PubMed]
  51. Fuchs, T.; Huber, H.; Burggraf, S.; Stetter, K.O. 16S rDNA-based Phylogeny of the Archaeal Order Sulfolobales and Reclassification of Desulfurolobus ambivalens as Acidianus ambivalens comb. nov. Syst. Appl. Microbiol. 1996, 19, 56–60. [Google Scholar] [CrossRef]
  52. Zillig, W.; Yeats, S.; Holz, I.; Böck, A.; Rettenberger, M.; Gropp, F.; Simon, G. Desulfurolobus ambivalens, gen. nov., sp. nov., an autotrophic archaebacterium facultatively oxidizing or reducing sulfur. Syst. Appl. Microbiol. 1986, 8, 197–203. [Google Scholar] [CrossRef]
  53. Brierley, C.L.; Brierley, J.A. A chemoautotrophic and thermophilic microorganism isolated from an acid hot spring. Can. J. Microbiol. 1973, 19, 183–188. [Google Scholar] [CrossRef] [PubMed]
  54. Konishi, Y.; Nishimura, H.; Asai, S. Bioleaching of sphalerite by the acidophilic thermophile Acidianus brierleyi. Hydrometallurgy 1998, 47, 339–352. [Google Scholar] [CrossRef]
  55. Dinkla, I.J.T.; Gericke, M.; Geurkink, B.K.; Hallberg, K.B. Acidianus brierleyi is the dominant thermoacidophile in a bioleaching community processing chalcopyrite containing concentrates at 70 °C. Adv. Mater. Res. 2009, 71, 67–70. [Google Scholar] [CrossRef]
  56. Brierley, J.A. Acdidophilic thermophilic archaebacteria: Potential application for metals recovery. FEMS Microbiol. Rev. 1990, 75, 287–292. [Google Scholar] [CrossRef]
  57. He, Z.G.; Zhong, H.; Li, Y. Acidianus tengchongensis sp. nov., a new species of acidothermophilic archaeon isolated from an Acidothermal Spring. Curr. Microbiol. 2004, 48, 159–163. [Google Scholar] [PubMed]
  58. Yoshida, N.; Nakasato, M.; Ohmura, N.; Ando, A.; Saiki, H.; Ishii, M.; Igarashi, Y. Acidianus manzaensis sp. nov., a novel thermoacidophilic Archaeon growing autotrophically by the oxidation of H2 with the reduction of Fe3+. Curr. Microbiol. 2006, 53, 406–411. [Google Scholar] [CrossRef] [PubMed]
  59. Ding, J.; Zhang, R.; Yu, Y.; Jin, D.; Liang, C.; Yi, Y.; Zhu, W.; Xia, J. A novel acidophilic, thermophilic iron and sulfur-oxidizing archaeon isolated from a hot spring of tengchong, Yunnan, China. Brazilian J. Microbiol. 2011, 42, 514–525. [Google Scholar] [CrossRef]
  60. Kurosawa, N.; Itoh, Y.H.; Iwai, T.; Sugai, A.; Uda, I.; Kimura, N.; Horiuchi, T.; Itoh, T. Sulfurisphaera ohwakuensis gen. nov., sp. nov., a novel extremely thermophilic acidophile of the order Sulfolobales. Int. J. Syst. Bacteriol. 1998, 48, 451–456. [Google Scholar] [CrossRef] [PubMed]
  61. Segerer, A.H.; Trincone, A.; Gahrtz, M.; Stetter, K.O. Stygiolobus azoricus gen. nov., sp. nov. represents a novel genus of anaerobic, extremely thermoacidophilic archaebacteria of the order sulfolobales. Int. J. Syst. Bacteriol. 1991, 41, 495–501. [Google Scholar] [CrossRef]
  62. Huber, H.; Prangishvili, D. Sulfolobales. In The Prokaryotes; Dworkin, M., Falkow, S., Rosenberg, E., Schleifer, K.-H., Stackebrant, E., Eds.; Springer Science: New York, NY, USA, 2006; pp. 1028–1049. [Google Scholar]
  63. Albers, S.-V.; Siebers, B. The family sulfolobaceae. In The Prokaryotes; Eugene, R., DeLong, E.F., Lory, S., Stackebrandt, E., Thompson, F., Eds.; Springer: Berlin, Germany, 2014; pp. 323–346. [Google Scholar]
  64. Rawlings, D.E.; Johnson, D.B. The microbiology of biomining: Development and optimization of mineral-oxidizing microbial consortia. Microbiology 2007, 153, 315–324. [Google Scholar] [CrossRef] [PubMed]
  65. Albers, S.-V.; Jarrell, K.F. The archaellum: How archaea swim. Front. Microbiol. 2015, 6, 1–12. [Google Scholar] [CrossRef] [PubMed]
  66. Mao, D.; Grogan, D. Genomic evidence of rapid, global-scale gene flow in a Sulfolobus species. ISME J. 2012, 6, 1613–1616. [Google Scholar] [CrossRef] [PubMed]
  67. Inskeep, W.P.; Jay, Z.J.; Tringe, S.G.; Herrgård, M.J.; Rusch, D.B. The YNP metagenome project: Environmental parameters responsible for microbial distribution in the yellowstone geothermal ecosystem. Front. Microbiol. 2013, 4, 1–15. [Google Scholar] [CrossRef] [PubMed]
  68. Reno, M.L.; Held, N.L.; Fields, C.J.; Burke, P.V.; Whitaker, R.J. Biogeography of the Sulfolobus islandicus pan-genome. Proc. Natl. Acad. Sci. USA 2009, 106, 8605–8610. [Google Scholar] [CrossRef] [PubMed]
  69. Guo, L.; Brügger, K.; Liu, C.; Shah, S.A.; Zheng, H.; Zhu, Y.; Wang, S.; Lillestøl, R.K.; Chen, L.; Frank, J.; et al. Genome analyses of icelandic strains of Sulfolobus islandicus, model organisms for genetic and virus-host interaction studies. J. Bacteriol. 2011, 193, 1672–1680. [Google Scholar] [CrossRef] [PubMed]
  70. Cadillo-Quiroz, H.; Didelot, X.; Held, N.L.; Herrera, A.; Darling, A.; Reno, M.L.; Krause, D.J.; Whitaker, R.J. Patterns of gene flow define species of thermophilic Archaea. PLoS Biol. 2012, 10. [Google Scholar] [CrossRef] [PubMed]
  71. Jaubert, C.; Danioux, C.; Oberto, J.; Cortez, D.; Bize, A.; Krupovic, M.; She, Q.; Forterre, P.; Prangishvili, D.; Sezonov, G. Genomics and genetics of Sulfolobus islandicus LAL14/1, a model hyperthermophilic archaeon. Open Biol. 2013, 3, 130010. [Google Scholar] [CrossRef] [PubMed]
  72. Wagner, M.; van Wolferen, M.; Wagner, A.; Lassak, K.; Meyer, B.H.; Reimann, J.; Albers, S.-V. Versatile genetic tool box for the crenarchaeote Sulfolobus acidocaldarius. Front. Microbiol. 2012, 3. [Google Scholar] [CrossRef] [PubMed]
  73. Albers, S.V.; Driessen, A.J.M. Conditions for gene disruption by homologous recombination of exogenous DNA into the Sulfolobus solfataricus genome. Archaea 2008, 2, 145–149. [Google Scholar] [CrossRef] [PubMed]
  74. Maezato, Y.; Dana, K.; Blum, P. Engineering thermoacidophilic archaea using linear DNA recombination. In Strain Engineering; Williams, J.A., Ed.; Humana Press: New York, NY, USA, 2011; Volume 765, pp. 435–445. [Google Scholar]
  75. Deng, L.; Zhu, H.; Chen, Z.; Liang, Y.X.; She, Q. Unmarked gene deletion and host-vector system for the hyperthermophilic crenarchaeon Sulfolobus islandicus. Extremophiles 2009, 13, 735–746. [Google Scholar] [CrossRef] [PubMed]
  76. Zhang, C.; Whitaker, R.J. A broadly applicable gene knockout system for the thermoacidophilic archaeon Sulfolobus islandicus based on simvastatin selection. Microbiology 2012, 158, 1513–1522. [Google Scholar] [CrossRef] [PubMed]
  77. Berkner, S.; Wlodkowski, A.; Albers, S.V.; Lipps, G. Inducible and constitutive promoters for genetic systems in Sulfolobus acidocaldarius. Extremophiles 2010, 14, 249–259. [Google Scholar] [CrossRef] [PubMed]
  78. Berkner, S.; Grogan, D.; Albers, S.V.; Lipps, G. Small multicopy, non-integrative shuttle vectors based on the plasmid pRN1 for Sulfolobus acidocaldarius and Sulfolobus solfataricus, model organisms of the (cren-)archaea. Nucl. Acids Res. 2007, 35, 1–12. [Google Scholar] [CrossRef] [PubMed]
  79. Zhang, C.; Krause, D.J.; Whitaker, R.J. Sulfolobus islandicus: A model system for evolutionary genomics. Biochem. Soc. Trans. 2013, 41, 458–462. [Google Scholar] [CrossRef] [PubMed]
  80. Zheng, T.; Huang, Q.; Zhang, C.; Ni, J.; She, Q.; Shen, Y. Development of a simvastatin selection marker for a hyperthermophilic acidophile, Sulfolobus islandicus. Appl. Environ. Microbiol. 2012, 78, 568–574. [Google Scholar] [CrossRef] [PubMed]
  81. López-García, P.; Forterre, P. Control of DNA topology during thermal stress in hyperthermophilic archaea: DNA topoisomerase levels, activities and induced thermotolerance during heat and cold shock in Sulfolobus. Mol. Microbiol. 1999, 33, 766–777. [Google Scholar] [CrossRef] [PubMed]
  82. Zillig, W.; Arnold, H.P.; Holz, I.; Prangishvili, D.; Schweier, A.; Stedman, K.; She, Q.; Phan, H.; Garrett, R.; Kristjansson, J.K. Genetic elements in the extremely thermophilic archaeon Sulfolobus. Extremophiles 1998, 2, 131–140. [Google Scholar] [CrossRef] [PubMed]
  83. Whitaker, R.J.; Grogan, D.W.; Taylor, J.W. Geographic barriers isolate endemic populations of hyperthermophilic archaea. Science 2003, 301, 976–978. [Google Scholar] [CrossRef] [PubMed]
  84. Grogan, D.W.; Ozarzak, M.A.; Bernander, R. Variation in gene content among geographically diverse Sulfolobus isolates. Environ. Microbiol. 2008, 10, 137–146. [Google Scholar] [CrossRef] [PubMed]
  85. Held, N.L.; Herrera, A.; Whitaker, R.J. Reassortment of CRISPR repeat-spacer loci in Sulfolobus islandicus. Environ. Microbiol. 2013, 15, 3065–3076. [Google Scholar] [PubMed]
  86. Bautista, M.A.; Zhang, C.; Whitaker, R.J. Virus-Induced Dormancy in the Archaeon Sulfolobus islandicus. MBio 2015, 6, e02565-14. [Google Scholar] [CrossRef] [PubMed]
  87. Peeples, T.L.; Kelly, R.M. Bioenergetics of the metal/sulfur-oxidizing extreme thermoacidophile, Metallosphaera sedula. Fuel 1993, 72, 1619–1624. [Google Scholar] [CrossRef]
  88. Berg, I.A.; Kockelkorn, D.; Buckel, W.; Fuchs, G. A 3-Hydroxypropionate/4-Hydroxybutyrate Autotrophic Carbon Dioxide Assimilation Pathway in Archaea. Science 2007, 318, 1782–1786. [Google Scholar] [CrossRef] [PubMed]
  89. Laska, S.; Lottspeich, F.; Kletzin, A. Membrane-bound hydrogenase and sulfur reductase of the hyperthermophilic and acidophilic archaeon Acidianus ambivalens. Microbiology 2003, 149, 2357–2371. [Google Scholar] [CrossRef] [PubMed]
  90. Podar, M.; Makarova, K.S.; Graham, D.E.; Wolf, Y.I.; Koonin, E.V.; Reysenbach, A.-L. Insights into archaeal evolution and symbiosis from the genomes of a nanoarchaeon and its inferred crenarchaeal host from Obsidian Pool, Yellowstone National Park. Biol. Direct 2013, 8, 9. [Google Scholar] [CrossRef] [PubMed]
  91. Servín-Garcidueñas, L.E.; Martínez-Romero, E. Draft Genome Sequence of the Sulfolobales Archaeon AZ1, Obtained through Metagenomic Analysis of a Mexican Hot Spring. Genome Announc. 2014, 2, 1–2. [Google Scholar] [CrossRef] [PubMed]
  92. Ingledew, W.J. Thiobacillus Ferrooxidans the bioenergetics of an acidophilic chemolithotroph. Biochim. Biophys. Acta 1982, 683, 89–117. [Google Scholar] [CrossRef]
  93. Bird, L.J.; Bonnefoy, V.; Newman, D.K. Bioenergetic challenges of microbial iron metabolisms. Trends Microbiol. 2011, 19, 330–340. [Google Scholar] [CrossRef] [PubMed]
  94. Bonnefoy, V.; Holmes, D.S. Genomic insights into microbial iron oxidation and iron uptake strategies in extremely acidic environments. Environ. Microbiol. 2012, 14, 1597–1611. [Google Scholar] [CrossRef] [PubMed]
  95. Bonnefoy, V. Bioinformatics and Genomics of Iron- and Sulfur-Oxidizing Acidophiles. In Geomicrobiology: Molecular and Environmental Perspective; Barton, L.L., Mandl, M., Loy, A., Eds.; Springer Netherlands: Dordrecht, The Netherlands, 2010; pp. 169–192. [Google Scholar]
  96. Quatrini, R.; Appia-Ayme, C.; Denis, Y.; Jedlicki, E.; Holmes, D.S.; Bonnefoy, V. Extending the models for iron and sulfur oxidation in the extreme acidophile Acidithiobacillus ferrooxidans. BMC Genomics 2009, 10, 394. [Google Scholar] [CrossRef] [PubMed]
  97. Blake, R.C.; Shute, E.A.; Greenwood, M.M.; Spencer, G.H.; Ingledew, W.J. Enzymes of aerobic respiration on iron. FEMS Microbiol. Rev. 1993, 11, 9–18. [Google Scholar] [CrossRef] [PubMed]
  98. Barr, D.W.; Ingledew, W.J.; Norris, P.R. Respiratory chain components of iron-oxidizing acidophilic bacteria. FEMS Microbiol. Lett. 1990, 70, 85–89. [Google Scholar] [CrossRef]
  99. Schäfer, G.; Engelhard, M.; Müller, V. Bioenergetics of the Archaea. Microbiol. Mol. Biol. Rev. 1999, 63, 570–620. [Google Scholar] [PubMed]
  100. Pereira, M.M.; Bandeiras, T.M.; Fernandes, A.S.; Lemos, R.S.; Melo, A.M.; Teixeira, M. Respiratory chains from aerobic thermophilic prokaryotes. J. Bioenerg. Biomembr. 2004, 36, 93–105. [Google Scholar] [CrossRef] [PubMed]
  101. Schmidt, C.L. Rieske Iron-Sulfur Proteins from Extremophilic Organisms. J. Bioenerg. Biomembr. 2004, 36, 107–113. [Google Scholar] [CrossRef] [PubMed]
  102. Lübben, M.; Kolmerer, B.; Saraste, M. An archaebacterial terminal oxidase combines core structures of two mitochondrial respiratory complexes. EMBO J. 1992, 11, 805–812. [Google Scholar] [PubMed]
  103. Lübben, M.; Warne, A.; Albracht, S.P.; Saraste, M. The purified SoxABCD quinol oxidase complex of Sulfolobus acidocaldarius contains a novel haem. Mol. Microbiol. 1994, 13, 327–335. [Google Scholar] [CrossRef] [PubMed]
  104. Gleißner, M.; Kaiser, U.; Antonopoulos, E.; Schäfer, G. The archaeal SoxABCD complex is a proton pump in Sulfolobus acidocaldarius. J. Biol. Chem. 1997, 272, 8417–8426. [Google Scholar] [CrossRef] [PubMed]
  105. Lübben, M.; Arnaud, S.; Castresana, J.; Warne, A.; Albracht, S.P.; Saraste, M. A second terminal oxidase in Sulfolobus acidocaldarius. Eur. J. Biochem. 1994, 224, 151–159. [Google Scholar] [CrossRef] [PubMed]
  106. Castresana, J.; Lübben, M.; Saraste, M. New archaebacterial genes coding for redox proteins: Implications for the evolution of aerobic metabolism. J. Mol. Biol. 1995, 250, 202–210. [Google Scholar] [CrossRef] [PubMed]
  107. Komorowski, L.; Schäfer, G. Sulfocyanin and subunit II, two copper proteins with novel features, provide new insight into the archaeal SoxM oxidase supercomplex. FEBS Lett. 2001, 487, 351–355. [Google Scholar] [CrossRef]
  108. Komorowski, L.; Verheyen, W.; Schäfer, G. The archaeal respiratory supercomplex SoxM from S. acidocaldarius combines features of quinole and cytochrome c oxidases. Biol. Chem. 2002, 383, 1791–1799. [Google Scholar] [CrossRef] [PubMed]
  109. Hiller, A.; Henninger, T.; Schäfer, G.; Schmidt, C.L. New genes encoding subunits of a cytochrome bc1-analogous complex in the respiratory chain of the hyperthermoacidophilic crenarchaeon Sulfolobus acidocaldarius. J. Bioenerg. Biomembr. 2003, 35, 121–131. [Google Scholar] [CrossRef] [PubMed]
  110. Bandeiras, T.M.; Refojo, P.N.; Todorovic, S.; Murgida, D.H.; Hildebrandt, P.; Bauer, C.; Pereira, M.M.; Kletzin, A.; Teixeira, M. The cytochrome ba complex from the thermoacidophilic crenarchaeote Acidianus ambivalens is an analog of bc(1) complexes. Biochim. Biophys. Acta 2009, 1787, 37–45. [Google Scholar] [CrossRef] [PubMed]
  111. Purschke, W.G.; Schmidt, C.L.; Petersen, A.; Schäfer, G. The terminal quinol oxidase of the hyperthermophilic archaeon Acidianus ambivalens exhibits a novel subunit structure and gene organization. J. Bacteriol. 1997, 179, 1344–1353. [Google Scholar] [PubMed]
  112. Giuffrè, A.; Gomes, C.M.; Antonini, G.; D’Itri, E.; Teixeira, M.; Brunori, M. Functional properties of the quinol oxidase from Acidianus ambivalens and the possible catalytic role of its electron donor—Studies on the membrane-integrated and purified enzyme. Eur. J. Biochem. 1997, 250, 383–388. [Google Scholar] [CrossRef] [PubMed]
  113. Das, T.K.; Gomes, C.M.; Bandeiras, T.M.; Pereira, M.M.; Teixeira, M.; Rousseau, D.L. Active site structure of the aa3 quinol oxidase of Acidianus ambivalens. Biochim. Biophys. Acta 2004, 1655, 306–320. [Google Scholar] [CrossRef] [PubMed]
  114. Kappler, U.; Sly, L.I.; McEwan, A.G. Respiratory gene clusters of Metallosphaera sedula—Differential expression and transcriptional organization. Microbiology 2005, 151, 35–43. [Google Scholar] [CrossRef] [PubMed]
  115. Hettmann, T.; Schmidt, C.L.; Anemüller, S.; Zähringer, U.; Moll, H.; Petersen, A.; Schäfer, G. Cytochrome b558/566 from the archaeon Sulfolobus acidocaldarius. A novel highly glycosylated, membrane-bound b-type hemoprotein. J. Biol. Chem. 1998, 273, 12032–12040. [Google Scholar] [CrossRef] [PubMed]
  116. Schoepp-Cothenet, B.; Schütz, M.; Baymann, F.; Brugna, M.; Nitschke, W.; Myllykallio, H.; Schmidt, C. The membrane-extrinsic domain of cytochrome b558/566 from the Archaeon Sulfolobus acidocaldarius performs pivoting movements with respect to the membrane surface. FEBS Lett. 2001, 487, 372–376. [Google Scholar] [CrossRef]
  117. Gomes, C.M.; Bandeiras, T.M.; Teixeira, M. A new type-II NADH dehydrogenase from the archaeon acidianus ambivalens: Characterization and in vitro reconstitution of the respiratory chain. J. Bioenerg. Biomembr. 2001, 33, 1–8. [Google Scholar] [CrossRef] [PubMed]
  118. Lemos, R.S.; Gomes, C.M.; Teixeira, M. Acidianus ambivalens Complex II typifies a novel family of succinate dehydrogenases. Biochem. Biophys. Res. Commun. 2001, 281, 141–150. [Google Scholar] [CrossRef] [PubMed]
  119. Iwasaki, T.; Kounosu, A.; Aoshima, M.; Ohmori, D.; Imai, T.; Urushiyama, A.; Cosper, N.J.; Scott, R.A. Novel [2Fe-2S]-type redox center C in sdhC of archaeal respiratory complex II from Sulfolobus tokodaii strain 7. J. Biol. Chem. 2002, 277, 39642–39648. [Google Scholar] [CrossRef] [PubMed]
  120. Kanao, T.; Kamimura, K.; Sugio, T. Identification of a gene encoding a tetrathionate hydrolase in Acidithiobacillus ferrooxidans. J. Biotechnol. 2007, 132, 16–22. [Google Scholar] [CrossRef] [PubMed]
  121. Bugaytsova, Z.; Lindström, E.B. Localization, purification and properties of a tetrathionate hydrolase from Acidithiobacillus caldus. Eur. J. Biochem. 2004, 271, 272–280. [Google Scholar] [CrossRef] [PubMed]
  122. Müller, F.H.; Bandeiras, T.M.; Urich, T.; Teixeira, M.; Gomes, C.M.; Kletzin, A. Coupling of the pathway of sulphur oxidation to dioxygen reduction: Characterization of a novel membrane-bound thiosulphate: Quinone oxidoreductase. Mol. Microbiol. 2004, 53, 1147–1160. [Google Scholar] [CrossRef] [PubMed]
  123. Castelle, C.; Guiral, M.; Malarte, G.; Ledgham, F.; Leroy, G.; Brugna, M.; Giudici-Orticoni, M.-T. A new iron-oxidizing/O2-reducing supercomplex spanning both inner and outer membranes, isolated from the extreme acidophile Acidithiobacillus ferrooxidans. J. Biol. Chem. 2008, 283, 25803–25811. [Google Scholar] [CrossRef] [PubMed]
  124. Yarzábal, A.; Brasseur, G.; Ratouchniak, J.; Lund, K.; Lemesle-Meunier, D.; DeMoss, J.A.; Bonnefoy, V. The high-molecular-weight cytochrome c Cyc2 of Acidithiobacillus ferrooxidans is an outer membrane protein. J. Bacteriol. 2002, 184, 313–317. [Google Scholar] [CrossRef] [PubMed]
  125. Bengrine, A.; Guiliani, N.; Appia-Ayme, C.; Jedlicki, E.; Holmes, D.S.; Chippaux, M.; Bonnefoy, V. Sequence and expression of the rusticyanin structural gene from Thiobacillus ferrooxidans ATCC33020 strain. Biochim. Biophys. Acta 1998, 1443, 99–112. [Google Scholar] [CrossRef]
  126. Giudici-Orticoni, M.T.; Guerlesquin, F.; Bruschi, M.; Nitschke, W. Interaction-induced redox switch in the electron transfer complex rusticyanin-cytochrome c(4). J. Biol. Chem. 1999, 274, 30365–30369. [Google Scholar] [CrossRef] [PubMed]
  127. Malarte, G.; Leroy, G.; Lojou, E.; Abergel, C.; Bruschi, M.; Giudici-Orticoni, M.T. Insight into molecular stability and physiological properties of the diheme cytochrome CYC41 from the acidophilic bacterium Acidithiobacillus ferrooxidans. Biochemistry 2005, 44, 6471–6481. [Google Scholar] [CrossRef] [PubMed]
  128. Kai, M.; Yano, T.; Tamegai, H.; Fukumori, Y.; Yamanaka, T. Thiobacillus ferrooxidans cytochrome c oxidase: Purification, and molecular and enzymatic features. J. Biochem. 1992, 112, 816–821. [Google Scholar] [PubMed]
  129. Appia-Ayme, C.; Guiliani, N.; Ratouchniak, J.; Bonnefoy, V. Characterization of an operon encoding two c-type cytochromes, an aa(3)-type cytochrome oxidase, and rusticyanin in Thiobacillus ferrooxidans ATCC 33020. Appl. Environ. Microbiol. 1999, 65, 4781–4787. [Google Scholar] [PubMed]
  130. Elbehti, A.; Brasseur, G.; Lemesle-Meunier, D. First evidence for existence of an uphill electron transfer through the bc(1) and NADH-Q oxidoreductase complexes of the acidophilic obligate chemolithotrophic ferrous ion-oxidizing bacterium Thiobacillus ferrooxidans. J. Bacteriol. 2000, 182, 3602–3606. [Google Scholar] [CrossRef] [PubMed]
  131. Brasseur, G.; Bruscella, P.; Bonnefoy, V.; Lemesle-Meunier, D. The bc1 complex of the iron-grown acidophilic chemolithotrophic bacterium Acidithiobacillus ferrooxidans functions in the reverse but not in the forward direction. Is there a second bc1 complex? Biochim. Biophys. Acta 2002, 1555, 37–43. [Google Scholar] [CrossRef]
  132. Brasseur, G.; Levican, G.; Bonnefoy, V.; Holmes, D.; Jedlicki, E.; Lemesle-Meunier, D. Apparent redundancy of electron transfer pathways via bc1 complexes and terminal oxidases in the extremophilic chemolithoautotrophic Acidithiobacillus ferrooxidans. Biochim. Biophys. Acta 2004, 1656, 114–126. [Google Scholar] [CrossRef] [PubMed]
  133. Dispirito, A.A.; Tuovinen, O.H. Uranous ion oxidation and carbon dioxide fixation by Thiobacillus ferrooxidans. Arch. Microbiol. 1982, 133, 28–32. [Google Scholar] [CrossRef]
  134. Finneran, K.T.; Housewright, M.E.; Lovley, D.R. Multiple influences of nitrate on uranium solubility during bioremediation of uranium-contaminated subsurface sediments. Environ. Microbiol. 2002, 4, 510–516. [Google Scholar] [CrossRef] [PubMed]
  135. Beller, H.R. Anaerobic, nitrate-dependent oxidation of U(IV) oxide minerals by the chemolithoautotrophic bacterium Thiobacillus denitrificans. Appl. Environ. Microbiol. 2005, 71, 2170–2174. [Google Scholar] [CrossRef] [PubMed]
  136. Beller, H.R.; Legler, T.C.; Bourguet, F.; Letain, T.E.; Kane, S.R.; Coleman, M. A Identification of c-type cytochromes involved in anaerobic, bacterial U(IV) oxidation. Biodegradation 2009, 20, 45–53. [Google Scholar] [CrossRef] [PubMed]
  137. Dispirito, A.A.; Tuovinen, O.H. Kinetics of uranous ion and ferrous iron oxidation by Thiobacillus ferrooxidans. Arch. Microbiol. 1982, 133, 33–37. [Google Scholar] [CrossRef]
  138. Nies, D.; Silver, S. Molecular Microbiology of Heavy Metals; Springer-Verlag: Berlin, Germany, 2007. [Google Scholar]
  139. Bruins, M.R.; Kapil, S.; Oehme, F.W. Microbial resistance to metals in the environment. Ecotoxicol. Environ. Saf. 2000, 45, 198–207. [Google Scholar] [CrossRef] [PubMed]
  140. Nies, D.H.; Silver, S. Ion efflux systems involved in bacterial metal resistances. J. Ind. Microbiol. 1995, 14, 186–199. [Google Scholar] [CrossRef] [PubMed]
  141. Dopson, M.; Holmes, D.S. Metal resistance in acidophilic microorganisms and its significance for biotechnologies. Appl. Microbiol. Biotechnol. 2014, 98, 8133–8144. [Google Scholar] [CrossRef] [PubMed]
  142. Navarro, C.A.; von Bernath, D.; Jerez, C.A. Heavy metal resistance strategies of acidophilic bacteria and their acquisition: Importance for biomining and bioremediation. Biol. Res. 2013, 46, 363–371. [Google Scholar] [CrossRef] [PubMed]
  143. Silver, S.; Phung, L.T. A bacterial view of the periodic table: Genes and proteins for toxic inorganic ions. J. Ind. Microbiol. Biotechnol. 2005, 32, 587–605. [Google Scholar] [CrossRef] [PubMed]
  144. Nies, D.H. Microbial heavy-metal resistance. Appl. Microbiol. Biotechnol. 1999, 51, 730–750. [Google Scholar] [CrossRef] [PubMed]
  145. Nies, D.H. Efflux-mediated heavy metal resistance in prokaryotes. FEMS Microbiol. Rev. 2003, 27, 313–339. [Google Scholar] [CrossRef]
  146. Orell, A.; Navarro, C.A.; Arancibia, R.; Mobarec, J.C.; Jerez, C.A. Life in blue: Copper resistance mechanisms of bacteria and archaea used in industrial biomining of minerals. Biotechnol. Adv. 2010, 28, 839–848. [Google Scholar] [CrossRef] [PubMed]
  147. Dopson, M.; Ossandon, F.J.; Lövgren, L.; Holmes, D.S. Metal resistance or tolerance? Acidophiles confront high metal loads via both abiotic and biotic mechanisms. Front. Microbiol. 2014, 5, 157. [Google Scholar] [CrossRef] [PubMed]
  148. Di Toro, D.M.; Allen, H.E.; Bergman, H.L.; Meyer, J.S.; Paquin, P.R.; Santore, R.C. Biotic ligand model of the acute toxicity of metals. 1. Technical basis. Environ. Toxicol. Chem. 2001, 20, 2383–2396. [Google Scholar] [CrossRef] [PubMed]
  149. Moberly, J.G.; Staven, A.R.I.; Sani, R.K. Influence of pH and inorganic phosphate on toxicity of zinc to Arthrobacter sp. isolated from sediments. Environ. Sci. Technol. 2010, 44, 7302–7308. [Google Scholar] [CrossRef] [PubMed]
  150. Baker-Austin, C.; Dopson, M. Life in acid: pH homeostasis in acidophiles. Trends Microbiol. 2007, 15, 165–171. [Google Scholar] [CrossRef] [PubMed]
  151. Slonczewski, J.L.; Fujisawa, M.; Dopson, M.; Krulwich, T.A. Cytoplasmic pH Measurement and Homeostasis in Bacteria and Archaea. Adv. Microb Physiol. 2009, 55, 1–79. [Google Scholar] [PubMed]
  152. Heijerick, D.G.; de Schamphelaere, K.A.C.; Janssen, C.R. Biotic ligand model development predicting Zn toxicity to the alga Pseudokirchneriella subcapitata: Possibilities and limitations. Comp. Biochem. Physiol. 2002, 133, 207–218. [Google Scholar] [CrossRef]
  153. Mertens, J.; Degryse, F.; Springael, D.; Smolders, E. Zinc toxicity to nitrification in soil and soilless culture can be predicted with the same biotic ligand model. Environ. Sci. Technol. 2007, 41, 2992–2997. [Google Scholar] [CrossRef] [PubMed]
  154. Mangold, S.; Potrykus, J.; Björn, E.; Lövgren, L.; Dopson, M. Extreme zinc tolerance in acidophilic microorganisms from the bacterial and archaeal domains. Extremophiles 2013, 75–85. [Google Scholar] [CrossRef] [PubMed]
  155. Maezato, Y.; Johnson, T.; McCarthy, S.; Dana, K.; Blum, P. Metal Resistance and Lithoautotrophy in the Extreme Thermoacidophile Metallosphaera Sedula. J. Bacteriol. 2012, 194, 6856–6863. [Google Scholar] [CrossRef] [PubMed]
  156. Cervantes, C.; Ji, G.; Ramírez, J.L.; Silver, S. Resistance to arsenic compounds in microorganisms. FEMS Microbiol. Rev. 1994, 15, 355–367. [Google Scholar] [CrossRef] [PubMed]
  157. McCarthy, S.; Ai, C.; Wheaton, G.; Tevatia, R.; Eckrich, V.; Kelly, R.; Blum, P. Role of an archaeal PitA transporter in the copper and arsenic resistance of Metallosphaera sedula, an extreme thermoacidophile. J. Bacteriol. 2014, 196, 3562–35670. [Google Scholar] [CrossRef] [PubMed]
  158. Bennett, R.L.; Malamy, M.H. Arsenate resistant mutants of Escherichia coli and phosphate transport. Biochem. Biophys. Res. Commun. 1970, 40, 496–503. [Google Scholar] [CrossRef]
  159. Willsky, G.R.; Malamy, M.H. Effect of arsenate on inorganic phosphate transport in Escherichia coli. J. Bacteriol. 1980, 144, 356–365. [Google Scholar] [PubMed]
  160. Nelson, D.L.; Kennedy, E.P. Magnesium transport in Escherichia coli. Inhibition by cobaltous ion. J. Biol. Chem. 1971, 246, 3042–3049. [Google Scholar] [PubMed]
  161. Park, M.H.; Wong, B.B.; Lusk, J.E. Mutants in three genes affecting transport of magnesium in Escherichia coli: Genetics and physiology. J. Bacteriol. 1976, 126, 1096–1103. [Google Scholar] [PubMed]
  162. Bini, E. Archaeal transformation of metals in the environment. FEMS Microbiol. Ecol. 2010, 73, 1–16. [Google Scholar] [CrossRef] [PubMed]
  163. Chivers, P.T.; Sauer, R.T. Regulation of high affinity nickel uptake in bacteria. Ni2+-dependent interaction of NikR with wild-type and mutant operator sites. J. Biol. Chem. 2000, 275, 19735–19741. [Google Scholar] [CrossRef] [PubMed]
  164. LaPaglia, C.; Hartzell, P.L. Stress-induced production of biofilm in thehyperthermophile Archaeoglobus fulgidus. Appl. Environ. Microbiol. 1997, 63, 3158–3163. [Google Scholar] [PubMed]
  165. Teitzel, G.M.; Parsek, M.R. Heavy Metal Resistance of Biofilm and Planktonic Pseudomonas aeruginosa Heavy Metal Resistance of Biofilm and Planktonic Pseudomonas aeruginosa. Appl. Environ. Microbiol. 2003, 69, 2313–2320. [Google Scholar] [CrossRef] [PubMed]
  166. Harrison, J.J.; Ceri, H.; Turner, R.J. Multimetal resistance and tolerance in microbial biofilms. Nat. Rev. Microbiol. 2007, 5, 928–938. [Google Scholar] [CrossRef] [PubMed]
  167. Macaskie, L.E.; Bonthrone, K.M.; Yong, P.; Goddard, D.T. Enzymically mediated bioprecipitation of uranium by a Citrobacter sp.: A concerted role for exocellular lipopolysaccharide and associated phosphatase in biomineral formation. Microbiology 2000, 146, 1855–1867. [Google Scholar] [PubMed]
  168. Merroun, M.; Hennig, C.; Rossberg, A.; Geipel, G.; Reich, T. Molecular and atomic analysis of uranium complexes formed by three eco-types of Acidithiobacillus ferrooxidans. Biochem. Soc. Trans. 2002, 30, 669–672. [Google Scholar] [CrossRef] [PubMed]
  169. Merroun, M.L.; Raff, J.; Rossberg, A.; Hennig, C.; Reich, T.; Selenska-pobell, S. Complexation of Uranium by Cells and S-Layer Sheets of Bacillus sphaericus JG-A12. Appl. Environ. Microbiol. 2005, 71, 5532–5543. [Google Scholar] [CrossRef] [PubMed]
  170. Sleytr, U.B.; Schuster, B.; Egelseer, E.M.; Pum, D. S-layers: Principles and applications. FEMS Microbiol. Rev. 2014, 38, 823–864. [Google Scholar] [CrossRef] [PubMed]
  171. Palmgren, M.G.; Nissen, P. P-type ATPases. Annu. Rev. Biophys. 2011, 40, 243–266. [Google Scholar] [CrossRef] [PubMed]
  172. Argüello, J.M. Identification of Ion-Selectivity Determinants in Heavy-Metal Transport P1B-type ATPases. J. Membr. Biol. 2003, 195, 93–108. [Google Scholar] [CrossRef] [PubMed]
  173. Argüello, J.M.; Eren, E.; González-Guerrero, M. The structure and function of heavy metal Transport P1B-ATPases. Biometals 2007, 20, 233–248. [Google Scholar] [CrossRef] [PubMed]
  174. Argüello, J.M.; González-Guerrero, M.; Raimunda, D. Bacterial transition metal P(1B)-ATPases: Transport mechanism and roles in virulence. Biochemistry 2011, 50, 9940–9949. [Google Scholar] [CrossRef] [PubMed]
  175. Moraleda-Muñoz, A.; Pérez, J.; Extremera, A.L.; Muñoz-Dorado, J. Expression and physiological role of three Myxococcus xanthus copper-dependent P1B-type ATPases during bacterial growth and development. Appl. Environ. Microbiol. 2010, 76, 6077–6084. [Google Scholar] [CrossRef] [PubMed]
  176. Rensing, C.; Ghosh, M.; Rosen, B.P. Families of soft-metal-ion-transporting ATPases. J. Bacteriol. 1999, 181, 5891–5897. [Google Scholar] [PubMed]
  177. Solioz, M.; Vulpe, C. CPx-type ATPases: A class of P-type ATPases that pump heavy metals. Trends Biochem. Sci. 1996, 21, 237–241. [Google Scholar] [CrossRef]
  178. Völlmecke, C.; Drees, S.L.; Reimann, J.; Albers, S.-V.; Lübben, M. The ATPases CopA and CopB both contribute to copper resistance of the thermoacidophilic archaeon Sulfolobus solfataricus. Microbiology 2012, 158, 1622–1633. [Google Scholar] [CrossRef] [PubMed]
  179. Chávez, F.P.; Lünsdorf, H.; Jerez, C.A. Growth of polychlorinated-biphenyl-degrading bacteria in the presence of biphenyl and chlorobiphenyls generates oxidative stress and massive accumulation of inorganic polyphosphate. Appl. Environ. Microbiol. 2004, 70, 3064–3072. [Google Scholar] [CrossRef] [PubMed]
  180. Seufferheld, M.J.; Alvarez, H.M.; Farias, M.E. Role of polyphosphates in microbial adaptation to extreme environments. Appl. Environ. Microbiol. 2008, 74, 5867–5874. [Google Scholar] [CrossRef] [PubMed]
  181. Rao, N.N.; Gómez-García, M.R.; Kornberg, A. Inorganic polyphosphate: Essential for growth and survival. Annu. Rev. Biochem. 2009, 78, 605–647. [Google Scholar] [CrossRef] [PubMed]
  182. Keasling, J.D. Regulation of intracellular toxic metals and other cations by hydrolysis of polyphosphate. Ann. N. Y. Acad. Sci. 1997, 829, 242–249. [Google Scholar] [CrossRef] [PubMed]
  183. Alvarez, S.; Jerez, C.A. Copper ions stimulate polyphosphate degradation and phosphate efflux in Acidithiobacillus ferrooxidans. Appl. Environ. Microbiol. 2004, 70, 5177–5182. [Google Scholar] [CrossRef] [PubMed]
  184. Remonsellez, F.; Orell, A.; Jerez, C.A. Copper tolerance of the thermoacidophilic archaeon Sulfolobus metallicus: Possible role of polyphosphate metabolism. Microbiology 2006, 152, 59–66. [Google Scholar] [CrossRef] [PubMed]
  185. Schurig-Briccio, L.A.; Gennis, R.B. Characterization of the PIB-Type ATPases Present in Thermus thermophilus. J. Bacteriol. 2012, 194, 4107–4113. [Google Scholar] [CrossRef] [PubMed]
  186. González-Guerrero, M.; Raimunda, D.; Cheng, X.; Argüello, J.M. Distinct functional roles of homologous Cu+ efflux ATPases in Pseudomonas aeruginosa. Mol. Microbiol. 2010, 78, 1246–1258. [Google Scholar] [CrossRef] [PubMed]
  187. Koch, H.G.; Winterstein, C.; Saribas, A.S.; Alben, J.O.; Daldal, F. Roles of the ccoGHIS gene products in the biogenesis of the cbb(3)-type cytochrome c oxidase. J. Mol. Biol. 2000, 297, 49–65. [Google Scholar] [CrossRef] [PubMed]
  188. Hassani, B.K.; Astier, C.; Nitschke, W.; Ouchane, S. CtpA, a copper-translocating P-type ATPase involved in the biogenesis of multiple copper-requiring enzymes. J. Biol. Chem. 2010, 285, 19330–19337. [Google Scholar] [CrossRef] [PubMed]
  189. Villafane, A.; Voskoboynik, Y.; Ruhl, I.; Sannino, D.; Maezato, Y.; Blum, P.; Bini, E. CopR of Sulfolobus solfataricus represents a novel class of archaeal-specific copper-responsive activators of transcription. Microbiology 2011, 157, 2808–2817. [Google Scholar] [CrossRef] [PubMed]
  190. Ettema, T.J.G.; Huynen, M.A.; de Vos, W.M.; van der Oost, J. TRASH: A novel metal-binding domain predicted to be involved in heavy-metal sensing, trafficking and resistance. Trends Biochem. Sci. 2003, 28, 170–173. [Google Scholar] [CrossRef]
  191. Villafane, A.A.; Voskoboynik, Y.; Cuebas, M.; Ruhl, I.; Bini, E. Response to excess copper in the hyperthermophile Sulfolobus solfataricus strain 98/2. Biochem. Biophys. Res. Commun. 2009, 385, 67–71. [Google Scholar] [CrossRef] [PubMed]
  192. Ettema, T.J.G.; Brinkman, A.B.; Lamers, P.P.; Kornet, N.G.; de Vos, W.M.; van der Oost, J. Molecular characterization of a conserved archaeal copper resistance (cop) gene cluster and its copper-responsive regulator in Sulfolobus solfataricus P2. Microbiology 2006, 152, 1969–1979. [Google Scholar] [CrossRef] [PubMed]
  193. Deigweiher, K.; Drell, T.L.; Prutsch, A.; Scheidig, A.J.; Lübben, M. Expression, isolation, and crystallization of the catalytic domain of CopB, a putative copper transporting ATPase from the thermoacidophilic archaeon Sulfolobus solfataricus. J. Bioenerg. Biomembr. 2004, 36, 151–159. [Google Scholar] [CrossRef] [PubMed]
  194. Mana-Capelli, S.; Mandal, A.K.; Argüello, J.M. Archaeoglobus fulgidus CopB is a thermophilic Cu2+-ATPase: Functional role of its histidine-rich-N-terminal metal binding domain. J. Biol. Chem. 2003, 278, 40534–40541. [Google Scholar] [CrossRef] [PubMed]
  195. Mandal, A.K.; Cheung, W.D.; Argüello, J.M. Characterization of a thermophilic P-type Ag+/Cu+-ATPase from the extremophile Archaeoglobus fulgidus. J. Biol. Chem. 2002, 277, 7201–7208. [Google Scholar] [CrossRef] [PubMed]
  196. Mandal, A.K.; Argüello, J.M. Functional roles of metal binding domains of the Archaeoglobus fulgidus Cu(+)-ATPase CopA. Biochemistry 2003, 42, 11040–11047. [Google Scholar] [CrossRef] [PubMed]
  197. Sazinsky, M.H.; Agarwal, S.; Argüello, J.M.; Rosenzweig, A.C. Structure of the actuator domain from the Archaeoglobus fulgidus Cu(+)-ATPase. Biochemistry 2006, 45, 9949–9955. [Google Scholar] [CrossRef] [PubMed]
  198. Sazinsky, M.H.; Mandal, A.K.; Argüello, J.M.; Rosenzweig, A.C. Structure of the ATP binding domain from the Archaeoglobus fulgidus Cu+-ATPase. J. Biol. Chem. 2006, 281, 11161–11166. [Google Scholar] [CrossRef] [PubMed]
  199. Agarwal, S.; Hong, D.; Desai, N.K.; Sazinsky, M.H.; Argüello, J.M.; Rosenzweig, A.C. Structure and interactions of the C-terminal metal binding domain of Archaeoglobus fulgidus CopA. Proteins 2010, 78, 2450–2458. [Google Scholar] [CrossRef] [PubMed]
  200. Padilla-Benavides, T.; McCann, C.J.; Arguello, J.M. The mechanism of Cu+ transport ATPases: Interaction with Cu+ chaperones and the role of transient metal binding sites. J. Biol. Chem. 2012, 288, 69–78. [Google Scholar] [CrossRef] [PubMed]
  201. Allen, G.S.; Wu, C.-C.; Cardozo, T.; Stokes, D.L. The architecture of CopA from Archeaoglobus fulgidus studied by cryo-electron microscopy and computational docking. Structure 2011, 19, 1219–1232. [Google Scholar] [CrossRef] [PubMed]
  202. González-Guerrero, M.; Argüello, J.M. Mechanism of Cu+-transporting ATPases: Soluble Cu+ chaperones directly transfer Cu+ to transmembrane transport sites. Proc. Natl. Acad. Sci. USA 2008, 105, 5992–5997. [Google Scholar] [CrossRef] [PubMed]
  203. Baker-Austin, C.; Dopson, M.; Wexler, M.; Sawers, R.G.; Bond, P.L. Molecular insight into extreme copper resistance in the extremophilic archaeon “Ferroplasma acidarmanus” Fer1. Microbiology 2005, 151, 2637–2646. [Google Scholar] [CrossRef] [PubMed]
  204. Orell, A.; Remonsellez, F.; Arancibia, R.; Jerez, C. A Molecular characterization of copper and cadmium resistance determinants in the biomining thermoacidophilic archaeon Sulfolobus metallicus. Archaea 2013, 2013, 1–16. [Google Scholar] [CrossRef] [PubMed]
  205. Kornberg, A.; Rao, N.N.; Ault-Riché, D. Inorganic polyphosphate: A molecule of many functions. Annu. Rev. Biochem. 1999, 68, 89–125. [Google Scholar] [CrossRef] [PubMed]
  206. Orell, A.; Navarro, C.A.; Rivero, M.; Aguilar, J.S.; Jerez, C.A. Inorganic polyphosphates in extremophiles and their possible functions. Extremophiles 2012, 16, 573–583. [Google Scholar] [CrossRef] [PubMed]
  207. Navarro, C.A.; Orellana, L.H.; Mauriaca, C.; Jerez, C.A. Transcriptional and functional studies of Acidithiobacillus ferrooxidans genes related to survival in the presence of copper. Appl. Environ. Microbiol. 2009, 75, 6102–6109. [Google Scholar] [CrossRef] [PubMed]
  208. Rensing, C.; Grass, G. Escherichia coli mechanisms of copper homeostasis in a changing environment. FEMS Microbiol. Rev. 2003, 27, 197–213. [Google Scholar] [CrossRef]
  209. Cardona, S.T.; Chávez, F.P.; Jerez, C.A. The exopolyphosphatase gene from sulfolobus solfataricus: Characterization of the first gene found to be involved in polyphosphate metabolism in archaea. Appl. Environ. Microbiol. 2002, 68, 4812–4819. [Google Scholar] [CrossRef] [PubMed]
  210. Mathema, V.B.; Thakuri, B.C.; Sillanpää, M. Bacterial mer operon-mediated detoxification of mercurial compounds: A short review. Arch. Microbiol. 2011, 193, 837–844. [Google Scholar] [CrossRef] [PubMed]
  211. Boyd, E.S.; Barkay, T. The mercury resistance operon: From an origin in a geothermal environment to an efficient detoxification machine. Front. Microbiol. 2012, 3, 349. [Google Scholar] [CrossRef] [PubMed]
  212. Barkay, T.; Kritee, K.; Boyd, E.; Geesey, G. A thermophilic bacterial origin and subsequent constraints by redox, light and salinity on the evolution of the microbial mercuric reductase. Environ. Microbiol. 2010, 12, 2904–2917. [Google Scholar] [CrossRef] [PubMed]
  213. Barkay, T.; Miller, S.M.; Summers, A.O. Bacterial mercury resistance from atoms to ecosystems. FEMS Microbiol. Rev. 2003, 27, 355–384. [Google Scholar] [CrossRef]
  214. Simbahan, J.; Kurth, E.; Schelert, J.; Dillman, A.; Moriyama, E.; Jovanovich, S.; Blum, P. Community analysis of a mercury hot spring supports occurrence of domain-specific forms of mercuric reductase. Appl. Environ. Microbiol. 2005, 71, 8836–8845. [Google Scholar] [CrossRef] [PubMed]
  215. King, J.K.; Kostka, J.E.; Frischer, M.E.; Saunders, F.M. Sulfate-reducing bacteria methylate mercury at variable rates in pure culture and in marine sediments. Appl. Environ. Microbiol. 2000, 66, 2430–2437. [Google Scholar] [CrossRef] [PubMed]
  216. Hintelmann, H. Organomercurials. Their formation and pathways in the environment. Met. Ions Life Sci. 2010, 7, 365–401. [Google Scholar] [PubMed]
  217. Lin, C.; Yee, N.; Barkay, T. Microbial Transformations in the Mercury Cycle; Liu, G., Cai, Y., O’Driscoll, N., Eds.; John Wiley & Sons: Hoboken, NJ, USA, 2011. [Google Scholar]
  218. Pak, K.; Bartha, R. Mercury methylation by interspecies hydrogen and acetate transfer between sulfidogens and methanogens. Appl. Environ. Microbiol. 1998, 64, 1987–1990. [Google Scholar] [PubMed]
  219. Silver, S.; Phung, L.T. Bacterial heavy metal resistance: New surprises. Annu. Rev. Microbiol. 1996, 50, 753–789. [Google Scholar] [CrossRef] [PubMed]
  220. Osborn, A.M.; Bruce, K.D.; Strike, P.; Ritchie, D.A. Distribution, diversity and evolution of the bacterial mercury resistance (mer) operon. FEMS Microbiol. Rev. 2006, 19, 239–262. [Google Scholar] [CrossRef]
  221. Schelert, J.; Dixit, V.; Hoang, V.; Simbahan, J.; Drozda, M.; Blum, P. Occurrence and characterization of mercury resistance in the hyperthermophilic archaeon Sulfolobus solfataricus by use of gene disruption. J. Bacteriol. 2004, 186, 427–437. [Google Scholar] [CrossRef] [PubMed]
  222. Schelert, J.; Drozda, M.; Dixit, V.; Dillman, A.; Blum, P. Regulation of mercury resistance in the crenarchaeote Sulfolobus solfataricus. J. Bacteriol. 2006, 188, 7141–7150. [Google Scholar] [CrossRef] [PubMed]
  223. Freedman, Z.; Zhu, C.; Barkay, T. Mercury Resistance and Mercuric Reductase Activities and Expression among Chemotrophic Thermophilic Aquificae. Appl. Environ. Microbiol. 2012, 78, 6568–6575. [Google Scholar] [CrossRef] [PubMed]
  224. Wang, Y.; Freedman, Z.; Lu-Irving, P.; Kaletsky, R.; Barkay, T. An initial characterization of the mercury resistance (mer) system of the thermophilic bacterium Thermus thermophilus HB27. FEMS Microbiol. Ecol. 2009, 67, 118–129. [Google Scholar] [CrossRef] [PubMed]
  225. Sugio, T.; Komoda, T.; Okazaki, Y.; Takeda, Y.; Nakamura, S.; Takeuchi, F. Volatilization of Metal Mercury from Organomercurials by Highly Mercury-Resistant Acidithiobacillus ferrooxidans MON-1. Biosci. Biotechnol. Biochem. 2010, 74, 1007–1012. [Google Scholar] [CrossRef] [PubMed]
  226. Sugio, T.; Iwahori, K.; Takeuchi, F.; Negishi, A.; Maeda, T.; Kamimura, K. Cytochrome c oxidase purified from a mercury-resistant strain of Acidithiobacillus ferrooxidans volatilizes mercury. J. Biosci. Bioeng. 2001, 92, 44–49. [Google Scholar] [CrossRef]
  227. Páez-Espino, D.; Tamames, J.; de Lorenzo, V.; Cánovas, D. Microbial responses to environmental arsenic. Biometals 2009, 22, 117–130. [Google Scholar] [CrossRef] [PubMed]
  228. Rosen, B.P.; Liu, Z. Transport pathways for arsenic and selenium: A minireview. Environ. Int. 2009, 35, 512–515. [Google Scholar] [CrossRef] [PubMed]
  229. Mukhopadhyay, R.; Rosen, B.P.; Phung, L.T.; Silver, S. Microbial arsenic: From geocycles to genes and enzymes. FEMS Microbiol. Rev. 2002, 26, 311–325. [Google Scholar] [CrossRef] [PubMed]
  230. Xu, C.; Zhou, T.; Kuroda, M.; Rosen, B.P. Metalloid resistance mechanisms in prokaryotes. J. Biochem. 1998, 123, 16–23. [Google Scholar] [CrossRef] [PubMed]
  231. Mukhopadhyay, R.; Rosen, B.P. Arsenate reductases in prokaryotes and eukaryotes. Environ. Health Perspect. 2002, 110, 745–748. [Google Scholar] [CrossRef] [PubMed]
  232. Meng, Y.-L.; Liu, Z.; Rosen, B.P. As(III) and Sb(III) uptake by GlpF and efflux by ArsB in Escherichia coli. J. Biol. Chem. 2004, 279, 18334–18341. [Google Scholar] [CrossRef] [PubMed]
  233. Dey, S.; Rosen, B.P. Dual mode of energy coupling by the oxyanion-translocating ArsB protein. J. Bacteriol. 1995, 177, 385–389. [Google Scholar] [PubMed]
  234. Lin, Y.-F.; Walmsley, A.R.; Rosen, B.P. An arsenic metallochaperone for an arsenic detoxification pump. Proc. Natl. Acad. Sci. USA 2006, 103, 15617–15622. [Google Scholar] [CrossRef] [PubMed]
  235. Kotze, A.A.; Tuffin, I.M.; Deane, S.M.; Rawlings, D.E. Cloning and characterization of the chromosomal arsenic resistance genes from Acidithiobacillus caldus and enhanced arsenic resistance on conjugal transfer of ars genes located on transposon TnAtcArs. Microbiology 2006, 152, 3551–3560. [Google Scholar] [CrossRef] [PubMed]
  236. Wang, L.; Chen, S.; Xiao, X.; Huang, X.; You, D.; Zhou, X.; Deng, Z. arsRBOCT arsenic resistance system encoded by linear plasmid pHZ227 in Streptomyces sp. strain FR-008. Appl. Environ. Microbiol. 2006, 72, 3738–3742. [Google Scholar] [CrossRef] [PubMed]
  237. Sekine, M.; Tanikawa, S.; Omata, S.; Saito, M.; Fujisawa, T.; Tsukatani, N.; Tajima, T.; Sekigawa, T.; Kosugi, H.; Matsuo, Y.; et al. Sequence analysis of three plasmids harboured in Rhodococcus erythropolis strain PR4. Environ. Microbiol. 2006, 8, 334–346. [Google Scholar] [CrossRef] [PubMed]
  238. Achour-Rokbani, A.; Cordi, A.; Poupin, P.; Bauda, P.; Billard, P. Characterization of the ars gene cluster from extremely arsenic-resistant Microbacterium sp. strain A33. Appl. Environ. Microbiol. 2010, 76, 948–955. [Google Scholar] [CrossRef] [PubMed]
  239. Butcher, B.G.; Deane, S.M.; Rawlings, D.E. The chromosomal arsenic resistance genes of Thiobacillus ferrooxidans have an unusual arrangement and confer increased arsenic and antimony resistance to Escherichia coli. Appl. Environ. Microbiol. 2000, 66, 1826–1833. [Google Scholar] [CrossRef] [PubMed]
  240. López-Maury, L.; Florencio, F.J.; Reyes, J.C. Arsenic sensing and resistance system in the cyanobacterium Synechocystis sp. strain PCC 6803. J. Bacteriol. 2003, 185, 5363–5371. [Google Scholar] [CrossRef] [PubMed]
  241. Vorontsov, I.I.; Minasov, G.; Brunzelle, J.S.; Shuvalova, L.; Kiryukhina, O.; Collart, F.R.; Anderson, W.F. Crystal structure of an apo form of Shigella flexneri ArsH protein with an NADPH-dependent FMN reductase activity. Protein Sci. 2007, 16, 2483–2490. [Google Scholar] [CrossRef] [PubMed]
  242. Baker-Austin, C.; Dopson, M.; Wexler, M.; Sawers, R.G.; Stemmler, A.; Rosen, B.P.; Bond, P.L. Extreme arsenic resistance by the acidophilic archaeon “Ferroplasma acidarmanus” Fer1. Extremophiles 2007, 11, 425–434. [Google Scholar] [CrossRef] [PubMed]
  243. Gihring, T.M.; Bond, P.L.; Peters, S.C.; Banfield, J.F. Arsenic resistance in the archaeon “Ferroplasma acidarmanus”: New insights into the structure and evolution of the ars genes. Extremophiles 2003, 7, 123–130. [Google Scholar] [PubMed]
  244. Ruepp, A.; Graml, W.; Santos-Martinez, M.L.; Koretke, K.K.; Volker, C.; Mewes, H.W.; Frishman, D.; Stocker, S.; Lupas, A.N.; Baumeister, W. The genome sequence of the thermoacidophilic scavenger Thermoplasma acidophilum. Nature 2000, 407, 508–513. [Google Scholar] [PubMed]
  245. Fütterer, O.; Angelov, A.; Liesegang, H.; Gottschalk, G.; Schleper, C.; Schepers, B.; Dock, C.; Antranikian, G.; Liebl, W. Genome sequence of Picrophilus torridus and its implications for life around pH 0. Proc. Natl. Acad. Sci. USA 2004, 101, 9091–9096. [Google Scholar] [CrossRef] [PubMed]
  246. Cozen, A.E.; Weirauch, M.T.; Pollard, K.S.; Bernick, D.L.; Stuart, J.M.; Lowe, T.M. Transcriptional map of respiratory versatility in the hyperthermophilic crenarchaeon Pyrobaculum aerophilum. J. Bacteriol. 2009, 191, 782–794. [Google Scholar] [CrossRef] [PubMed]
  247. Dopson, M.; Lindström, E.B.; Hallberg, K.B. Chromosomally encoded arsenical resistance of the moderately thermophilic acidophile Acidithiobacillus caldus. Extremophiles 2001, 5, 247–255. [Google Scholar] [CrossRef] [PubMed]
  248. Suzuki, K.; Wakao, N.; Kimura, T.; Sakka, K.; Ohmiya, K. Expression and regulation of the arsenic resistance operon of Acidiphilium multivorum AIU 301 plasmid pKW301 in Escherichia coli. Appl. Environ. Microbiol. 1998, 64, 411–418. [Google Scholar] [PubMed]
  249. Tuffin, I.M.; Hector, S.B.; Deane, S.M.; Rawlings, D.E. Resistance determinants of a highly arsenic-resistant strain of Leptospirillum ferriphilum isolated from a commercial biooxidation tank. Appl. Environ. Microbiol. 2006, 72, 2247–2253. [Google Scholar] [CrossRef] [PubMed]
  250. Wang, G.; Kennedy, S.P.; Fasiludeen, S.; Rensing, C.; DasSarma, S. Arsenic resistance in Halobacterium sp. strain NRC-1 examined by using an improved gene knockout system. J. Bacteriol. 2004, 186, 3187–3194. [Google Scholar] [CrossRef] [PubMed]
  251. Qin, J.; Rosen, B.P.; Zhang, Y.; Wang, G.; Franke, S.; Rensing, C. Arsenic detoxification and evolution of trimethylarsine gas by a microbial arsenite S-adenosylmethionine methyltransferase. Proc. Natl. Acad. Sci. USA 2006, 103, 2075–2080. [Google Scholar] [CrossRef] [PubMed]
  252. Cai, L.; Liu, G.; Rensing, C.; Wang, G. Genes involved in arsenic transformation and resistance associated with different levels of arsenic-contaminated soils. BMC Microbiol. 2009, 9, 4. [Google Scholar] [CrossRef] [PubMed]
  253. Sehlin, H.M.; Lindstrom, E.B. Oxidation and reduction of arsenic by Sulfolobus acidocaldarius strain BC. FEMS Microbiol. Lett. 1992, 93, 87–92. [Google Scholar] [CrossRef]
  254. Okibe, N.; Koga, M.; Sasaki, K.; Hirajima, T.; Heguri, S.; Asano, S. Simultaneous oxidation and immobilization of arsenite from refinery waste water by thermoacidophilic iron-oxidizing archaeon, Acidianus brierleyi. Miner. Eng. 2013, 48, 126–134. [Google Scholar] [CrossRef]
  255. Battaglia-Brunet, F.; Crouzet, C.; Breeze, D.; Tris, H.; Morin, D. Decreased leachability of arsenic linked to biological oxidation of As(III) in solid wastes from bioleaching liquors. Hydrometallurgy 2011, 107, 34–39. [Google Scholar] [CrossRef]
  256. Van Lis, R.; Nitschke, W.; Duval, S.; Schoepp-Cothenet, B. Arsenics as bioenergetic substrates. Biochim. Biophys. Acta 2013, 1827, 176–188. [Google Scholar] [CrossRef] [PubMed]
  257. Lebrun, E.; Brugna, M.; Baymann, F.; Muller, D.; Lièvremont, D.; Lett, M.-C.; Nitschke, W. Arsenite oxidase, an ancient bioenergetic enzyme. Mol. Biol. Evol. 2003, 20, 686–693. [Google Scholar] [CrossRef] [PubMed]
  258. Lett, M.-C.; Muller, D.; Lièvremont, D.; Silver, S.; Santini, J. Unified nomenclature for genes involved in prokaryotic aerobic arsenite oxidation. J. Bacteriol. 2012, 194, 207–208. [Google Scholar] [CrossRef] [PubMed]
  259. Dopson, M. Growth in sulfidic mineral environments: Metal resistance mechanisms in acidophilic micro-organisms. Microbiology 2003, 149, 1959–1970. [Google Scholar] [CrossRef] [PubMed]
  260. Salzano, A.M.; Febbraio, F.; Farias, T.; Cetrangolo, G.P.; Nucci, R.; Scaloni, A.; Manco, G. Redox stress proteins are involved in adaptation response of the hyperthermoacidophilic archaeon Sulfolobus solfataricus to nickel challenge. Microb. Cell Fact. 2007, 6, 25. [Google Scholar] [CrossRef] [PubMed]
  261. Lagorce, A.; Fourçans, A.; Dutertre, M.; Bouyssiere, B.; Zivanovic, Y.; Confalonieri, F. Genome-wide transcriptional response of the archaeon Thermococcus gammatolerans to cadmium. PLoS ONE 2012, 7, e41935. [Google Scholar] [CrossRef] [PubMed]
  262. Tian, J.; Wu, N.; Li, J.; Liu, Y.; Guo, J.; Yao, B.; Fan, Y. Nickel-resistant determinant from Leptospirillum ferriphilum. Appl. Environ. Microbiol. 2007, 73, 2364–2368. [Google Scholar] [CrossRef] [PubMed]
  263. Zhu, T.; Tian, J.; Zhang, S.; Wu, N.; Fan, Y. Identification of the transcriptional regulator NcrB in the nickel resistance determinant of Leptospirillum ferriphilum UBK03. PLoS ONE 2011, 6, e17367. [Google Scholar] [CrossRef] [PubMed]
  264. Merroun, M.L.; Selenska-Pobell, S. Bacterial interactions with uranium: An environmental perspective. J. Contam. Hydrol. 2008, 102, 285–295. [Google Scholar] [CrossRef] [PubMed]
  265. Cologgi, D.L.; Lampa-Pastirk, S.; Speers, A.M.; Kelly, S.D.; Reguera, G. Extracellular reduction of uranium via Geobacter conductive pili as a protective cellular mechanism. Proc. Natl. Acad. Sci. USA 2011, 108, 15248–15252. [Google Scholar] [CrossRef] [PubMed]
  266. Martinez, R.J.; Beazley, M.J.; Taillefert, M.; Arakaki, A.K.; Skolnick, J.; Sobecky, P.A. Aerobic uranium (VI) bioprecipitation by metal-resistant bacteria isolated from radionuclide- and metal-contaminated subsurface soils. Environ. Microbiol. 2007, 9, 3122–3133. [Google Scholar] [CrossRef] [PubMed]
  267. Renninger, N.; Knopp, R.; Nitsche, H.; Clark, D.S.; Keasling, J.D. Uranyl precipitation by Pseudomonas aeruginosa via controlled polyphosphate metabolism. Appl. Environ. Microbiol. 2004, 70, 7404–7412. [Google Scholar] [CrossRef] [PubMed]
  268. Merroun, M.L.; Nedelkova, M.; Ojeda, J.J.; Reitz, T.; Fernández, M.L.; Arias, J.M.; Romero-González, M.; Selenska-Pobell, S. Bio-precipitation of uranium by two bacterial isolates recovered from extreme environments as estimated by potentiometric titration, TEM and X-ray absorption spectroscopic analyses. J. Hazard. Mater. 2011, 197, 1–10. [Google Scholar] [CrossRef] [PubMed]
  269. Reitz, T.; Merroun, M.L.; Rossberg, A.; Steudtner, R.; Selenska-Pobell, S. Bioaccumulation of U(VI) by Sulfolobus acidocaldarius under moderate acidic conditions. Radiochim. Acta 2011, 99, 543–554. [Google Scholar] [CrossRef]
  270. Kashefi, K.; Lovley, D.R. Reduction of Fe(III), Mn(IV), and Toxic Metals at 100 C by Pyrobaculum islandicum. Appl. Environ. Microbiol. 2000, 66, 1050–1056. [Google Scholar] [CrossRef] [PubMed]
  271. Francis, A.J.; Gillow, J.B.; Dodge, C.J.; Harris, R.; Beveridge, T.J.; Papenguth, H.W. Uranium association with halophilic and non-halophilic bacteria and archaea. Radiochim. Acta 2004, 92, 481–488. [Google Scholar] [CrossRef]
  272. Reitz, T.; Merroun, M.; Rossberg, A.; Selenska-Pobell, S. Interactions of Sulfolobus acidocaldarius with uranium. Radiochim. Acta 2010, 98, 249–257. [Google Scholar] [CrossRef]
  273. Beazley, M.J.; Martinez, R.J.; Sobecky, P.A.; Webb, S.M.; Taillefert, M. Uranium biomineralization as a result of bacterial phosphatase activity: Insights from bacterial isolates from a contaminated subsurface. Environ. Sci. Technol. 2007, 41, 5701–5707. [Google Scholar] [CrossRef] [PubMed]
  274. Selenska-pobell, S.; Merroun, M. Accumulation of heavy metals by microorganisms: Bio-mineralization and nanocluster formation. In Prokaryotic Cell Wall Compounds; König, H., Claus, H., Varma, A., Eds.; Springer: Berlin, Germany, 2010; pp. 483–500. [Google Scholar]
  275. Merroun, M.; Hennig, C.; Rossberg, A.; Reich, T.; Selenska-Pobell, S. Characterization of U(VI)-Acidithiobacillus ferrooxidans complexes using EXAFS, transmission electron microscopy, and energy-dispersive X-ray analysis. Radiochim. Acta 2003, 91, 583–592. [Google Scholar] [CrossRef]
  276. Malekzadeh, F.; Latifi, A.M.; Shahamat, M.; Levin, M.; Colwell, R.R. Effects of selected physical and chemical parameters on uranium uptake by the bacterium Chryseomonas MGF-48. World J. Microbiol. Biotechnol. 2002, 18, 599–602. [Google Scholar] [CrossRef]
  277. Panak, P.J.; Knopp, R.; Booth, C.H.; Nitsche, H. Spectroscopic studies on the interaction of U(VI) with Bacillus sphaericus. Radiochim. Acta 2002, 90, 779–783. [Google Scholar] [CrossRef]
  278. Bäuerlein, E. Biomineralization of unicellular organisms: An unusual membrane biochemistry for the production of inorganic nano- and microstructures. Angew. Chem. Int. Ed. 2003, 42, 614–641. [Google Scholar] [CrossRef] [PubMed]
  279. Eichler, J. Facing extremes: Archaeal surface-layer (glyco)proteins. Microbiology 2003, 149, 3347–3351. [Google Scholar] [CrossRef] [PubMed]
  280. Dispirito, A.A.; Dugan, P.R.; Tuovinen, O.H. Sorption of Thiobacillus ferrooxidans to particulate material. Biotechnol. Bioeng. 1983, 25, 1163–1168. [Google Scholar] [CrossRef] [PubMed]
  281. Gehrke, T.; Telegdi, J.; Thierry, D.; Sand, W. Importance of Extracellular Polymeric Substances from Thiobacillus ferrooxidans for Bioleaching. Appl. Environ. Microbiol. 1998, 64, 2743–2747. [Google Scholar] [PubMed]
  282. Harneit, K.; Göksel, A.; Kock, D.; Klock, J.H.; Gehrke, T.; Sand, W. Adhesion to metal sulfide surfaces by cells of Acidithiobacillus ferrooxidans, Acidithiobacillus thiooxidans and Leptospirillum ferrooxidans. Hydrometallurgy 2006, 83, 245–254. [Google Scholar] [CrossRef]
  283. Africa, C.J.; van Hille, R.P.; Sand, W.; Harrison, S.T.L. Investigation and in situ visualisation of interfacial interactions of thermophilic microorganisms with metal-sulphides in a simulated heap environment. Miner. Eng. 2013, 48, 100–107. [Google Scholar] [CrossRef]
  284. Zhang, R.; Bellenberg, S.; Castro, L.; Neu, T.R.; Sand, W.; Vera, M. Colonization and biofilm formation of the extremely acidophilic archaeon Ferroplasma acidiphilum. Hydrometallurgy 2014, 150, 245–252. [Google Scholar] [CrossRef]
  285. Noël, N.; Florian, B.; Sand, W. AFM & EFM study on attachment of acidophilic leaching organisms. Hydrometallurgy 2010, 104, 370–375. [Google Scholar]
  286. Rodriguez-Leiva, M.; Tributsch, H. Morphology of bacterial leaching patterns by Thiobacillus ferrooxidans on synthetic pyrite. Arch. Microbiol. 1988, 149, 401–405. [Google Scholar] [CrossRef]
  287. Sand, W.; Gerke, T.; Hallmann, R.; Schippers, A. Sulfur chemistry, biofilm, and the (in)direct attack—A critical evaluation of bacterial leaching. Appl. Microbiol. Biotechnol. 1995, 43, 961–966. [Google Scholar] [CrossRef]
  288. Sand, W.; Gehrke, T.; Jozsa, P.G.; Schippers, A. (Bio)chemistry of bacterial leaching—Direct vs. indirect bioleaching. Hydrometallurgy 2001, 59, 159–175. [Google Scholar] [CrossRef]
  289. Sand, W.; Gehrke, T. Extracellular polymeric substances mediate bioleaching/biocorrosion via interfacial processes involving iron(III) ions and acidophilic bacteria. Res. Microbiol. 2006, 157, 49–56. [Google Scholar] [CrossRef] [PubMed]
  290. Florian, B.; Noël, N.; Thyssen, C.; Felschau, I.; Sand, W. Some quantitative data on bacterial attachment to pyrite. Miner. Eng. 2011, 24, 1132–1138. [Google Scholar] [CrossRef]
  291. Vera, M.; Schippers, A.; Sand, W. Progress in bioleaching: Fundamentals and mechanisms of bacterial metal sulfide oxidation-part A. Appl. Microbiol. Biotechnol. 2013, 97, 7529–7541. [Google Scholar] [CrossRef] [PubMed]
  292. Schippers, A.; Hedrich, S.; Vasters, J.; Drobe, M.; Sand, W.; Willscher, S. Biomining : Metal Recovery from Ores with Microorganisms. Adv. Biochem. Eng. Biotechnol. 2013, 141, 1–47. [Google Scholar]
  293. Vandevivere, P.; Kirchman, D.L. Attachment stimulates exopolysaccharide synthesis by a bacterium. Appl. Environ. Microbiol. 1993, 59, 3280–3286. [Google Scholar] [PubMed]
  294. Gehrke, T.; Hallmann, R.; Kinzler, K.; Sand, W. The EPS of Acidithiobacillus ferrooxidans—A model for structure-function relationships of attached bacteria and their physiology. Water Sci. Technol. 2001, 43, 159–167. [Google Scholar] [PubMed]
  295. Solari, J.A.; Huerta, G.; Escobar, B.; Vargas, T.; Badilla-Ohlbaum, R.; Rubio, J. Interfacial phenomena affecting the adhesion of Thiobacillus ferrooxidans to sulphide mineral surface. Colloids Surfaces 1992, 69, 159–166. [Google Scholar] [CrossRef]
  296. Blake, R.C.; Shute, E.A.; Howard, G.T. Solubilization of minerals by bacteria: Electrophoretic mobility of Thiobacillus ferrooxidans in the presence of iron, pyrite, and sulfur. Appl. Environ. Microbiol. 1994, 60, 3349–3357. [Google Scholar] [PubMed]
  297. Vilinska, A.; Rao, K.H. Surface Thermodynamics and Extended DLVO Theory of Acidithiobacillus ferrooxidans Cells Adhesion on Pyrite and Chalcopyrite. Open Colloid Sci. J. 2009, 2, 1–14. [Google Scholar] [CrossRef]
  298. Sampson, M.I.; Phillips, C.V.; Blake, R.C. Influence of the attachment of acidophilic bacteria during the oxidation of mineral sulfides. Miner. Eng. 2000, 13, 373–389. [Google Scholar] [CrossRef]
  299. Zhu, J.; Li, Q.; Jiao, W.; Jiang, H.; Sand, W.; Xia, J.; Liu, X.; Qin, W.; Qiu, G.; Hu, Y.; Chai, L. Adhesion forces between cells of Acidithiobacillus ferrooxidans, Acidithiobacillus thiooxidans or Leptospirillum ferrooxidans and chalcopyrite. Colloids Surfaces B Biointerfaces 2012, 94, 95–100. [Google Scholar] [CrossRef] [PubMed]
  300. Andrews, G.F. The selective adsorption of Thiobacilli to dislocation sites on pyrite surfaces. Biotechnol. Bioeng. 1988, 31, 378–381. [Google Scholar] [CrossRef] [PubMed]
  301. Edwards, K.J.; Rutenberg, A.D. Microbial response to surface microtopography: The role of metabolism in localized mineral dissolution. Chem. Geol. 2001, 180, 19–32. [Google Scholar] [CrossRef]
  302. Ohmura, N.; Kitamura, K.; Saiki, H. Selective adhesion of Thiobacillus ferrooxidans to pyrite. Appl. Environ. Microbiol. 1993, 59, 4044–4050. [Google Scholar] [PubMed]
  303. Sanhueza, A.; Ferrer, I.J.; Vargas, T.; Amils, R.; Sánchez, C. Attachment of Thiobacillus ferrooxidans on synthetic pyrite of varying structural and electronic properties. Hydrometallurgy 1999, 51, 115–129. [Google Scholar] [CrossRef]
  304. Shrihari; Modak, J.M.; Kumar, R.; Gandhi, K.S. Dissolution of particles of pyrite mineral by direct attachment of Thiobacillus ferrooxidans. Hydrometallurgy 1995, 38, 175–187. [Google Scholar] [CrossRef]
  305. Li, Y.-Q.; Wan, D.-S.; Huang, S.-S.; Leng, F.-F.; Yan, L.; Ni, Y.-Q.; Li, H.-Y. Type IV pili of Acidithiobacillus ferrooxidans are necessary for sliding, twitching motility, and adherence. Curr. Microbiol. 2010, 60, 17–24. [Google Scholar] [CrossRef] [PubMed]
  306. Pohlschroder, M.; Ghosh, A.; Tripepi, M.; Albers, S.-V. Archaeal type IV pilus-like structures—Evolutionarily conserved prokaryotic surface organelles. Curr. Opin. Microbiol. 2011, 14, 357–363. [Google Scholar] [CrossRef] [PubMed]
  307. Beveridge, T.J.; Pouwels, P.H.; Sára, M.; Kotiranta, A.; Lounatmaa, K.; Kari, K.; Kerosuo, E.; Haapasalo, M.; Egelseer, E.M.; Schocher, I.; et al. Functions of S-layers. FEMS Microbiol. Rev. 2006, 20, 99–149. [Google Scholar] [CrossRef]
  308. Acuña, J.; Rojas, J.; Amaro, A.M.; Toledo, H.; Jerez, C.A. Chemotaxis of Leptospirillum ferrooxidans and other acidophilic chemolithotrophs: Comparison with the Escherichia coli chemosensory system. FEMS Microbiol. Lett. 1992, 75, 37–42. [Google Scholar] [CrossRef]
  309. Meyer, G.; Schneider-Merk, T.; Böhme, S.; Sand, W. A simple method for investigations on the chemotaxis of Acidithiobacillus ferrooxidans and Desulfovibrio vulgaris. Acta Biotechnol. 2002, 22, 391–399. [Google Scholar] [CrossRef]
  310. Banderas, A.; Guiliani, N. Bioinformatic prediction of gene functions regulated by quorum sensing in the bioleaching bacterium Acidithiobacillus ferrooxidans. Int. J. Mol. Sci. 2013, 14, 16901–16916. [Google Scholar] [CrossRef] [PubMed]
  311. Bellenberg, S.; Díaz, M.; Noël, N.; Sand, W.; Poetsch, A.; Guiliani, N.; Vera, M. Biofilm formation, communication and interactions of leaching bacteria during colonization of pyrite and sulfur surfaces. Res. Microbiol. 2014, 165, 773–781. [Google Scholar] [CrossRef] [PubMed]
  312. Ruiz, L.M.; Castro, M.; Barriga, A.; Jerez, C.A.; Guiliani, N. The extremophile Acidithiobacillus ferrooxidans possesses a c-di-GMP signalling pathway that could play a significant role during bioleaching of minerals. Lett. Appl. Microbiol. 2012, 54, 133–139. [Google Scholar] [CrossRef] [PubMed]
  313. Castro, M.; Deane, S.M.; Ruiz, L.; Rawlings, D.E.; Guiliani, N. Diguanylate Cyclase Null Mutant Reveals that C-Di-GMP Pathway Regulates the Motility and Adherence of the Extremophile Bacterium Acidithiobacillus caldus. PLoS ONE 2015, 10, e0116399. [Google Scholar] [CrossRef] [PubMed]
  314. González, A.; Bellenberg, S.; Mamani, S.; Ruiz, L.; Echeverría, A.; Soulère, L.; Doutheau, A.; Demergasso, C.; Sand, W.; Queneau, Y.; et al. AHL signaling molecules with a large acyl chain enhance biofilm formation on sulfur and metal sulfides by the bioleaching bacterium Acidithiobacillus ferrooxidans. Appl. Microbiol. Biotechnol. 2013, 97, 3729–3737. [Google Scholar] [CrossRef] [PubMed]
  315. Farah, C.; Vera, M.; Morin, D.; Haras, D.; Jerez, C.A.; Guiliani, N. Evidence for a functional quorum-sensing type AI-1 system in the extremophilic bacterium Acidithiobacillus ferrooxidans. Appl. Environ. Microbiol. 2005, 71, 7033–7040. [Google Scholar] [CrossRef] [PubMed]
  316. Bellenberg, S.; Leon-Morales, C.-F.; Sand, W.; Vera, M. Visualization of capsular polysaccharide induction in Acidithiobacillus ferrooxidans. Hydrometallurgy 2012, 129, 82–89. [Google Scholar] [CrossRef]
  317. Vera, M.A.; Rohwerder, T.; Bellenberg, S.; Sand, W.; Denis, Y.; Bonnefoy, V. Characterization of biofilm formation by the bioleaching acidophilic bacterium acidithiobacillus ferrooxidans by a microarray transcriptome analysis. Adv. Mater. Res. 2009, 71, 175–178. [Google Scholar] [CrossRef]
  318. Barreto, M.; Jedlicki, E.; Holmes, D.S. Identification of a gene cluster for the formation of extracellular polysaccharide precursors in the chemolithoautotroph Acidithiobacillus ferrooxidans. Appl. Environ. Microbiol. 2005, 71, 2902–2909. [Google Scholar] [CrossRef] [PubMed]
  319. Moreno-Paz, M.; Gómez, M.J.; Arcas, A.; Parro, V. Environmental transcriptome analysis reveals physiological differences between biofilm and planktonic modes of life of the iron oxidizing bacteria Leptospirillum spp. in their natural microbial community. BMC Genomics 2010, 11, 404. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  320. Vera, M.; Krok, B.; Bellenberg, S.; Sand, W.; Poetsch, A. Shotgun proteomics study of early biofilm formation process of Acidithiobacillus ferrooxidans ATCC 23270 on pyrite. Proteomics 2013, 13, 1133–1144. [Google Scholar] [CrossRef] [PubMed]
  321. Dixon, D.G. Analysis of heat conservation during copper sulphide heap leaching. Hydrometallurgy 2000, 58, 27–41. [Google Scholar] [CrossRef]
  322. Olson, G.J.; Brierley, J.A.; Brierley, C.L. Bioleaching review part B: Progress in bioleaching: Applications of microbial processes by the minerals industries. Appl. Microbiol. Biotechnol. 2003, 63, 249–257. [Google Scholar] [CrossRef] [PubMed]
  323. Brierley, C.L.; Brierley, J.A. Progress in bioleaching: Part B: Applications of microbial processes by the minerals industries. Appl. Microbiol. Biotechnol. 2013, 97, 7543–7552. [Google Scholar] [CrossRef] [PubMed]
  324. Rodríguez, Y.; Ballester, A.; Blázquez, M.L.; González, F.; Muñoz, J.A. New information on the chalcopyrite bioleaching mechanism at low and high temperature. Hydrometallurgy 2003, 71, 47–56. [Google Scholar] [CrossRef]
  325. Mikkelsen, D.; Kappler, U.; Webb, R.I.; Rasch, R.; McEwan, A.G.; Sly, L.I. Visualisation of pyrite leaching by selected thermophilic archaea: Nature of microorganism-ore interactions during bioleaching. Hydrometallurgy 2007, 88, 143–153. [Google Scholar] [CrossRef]
  326. Gautier, V.; Escobar, B.; Vargas, T. Cooperative action of attached and planktonic cells during bioleaching of chalcopyrite with Sulfolobus metallicus at 70 °C. Hydrometallurgy 2008, 94, 121–126. [Google Scholar] [CrossRef]
  327. Bromfield, L.; Africa, C.J.; Harrison, S.T.L.; van Hille, R.P. The effect of temperature and culture history on the attachment of Metallosphaera hakonensis to mineral sulfides with application to heap bioleaching. Miner. Eng. 2011, 24, 1157–1165. [Google Scholar] [CrossRef]
  328. Zolghadr, B.; Kling, A.; Koerdt, A.; Driessen, A.J.M.; Rachel, R.; Albers, S.V. Appendage-mediated surface adherence of Sulfolobus solfataricus. J. Bacteriol. 2010, 192, 104–110. [Google Scholar] [CrossRef] [PubMed]
  329. Koerdt, A.; Orell, A.; Pham, T.K.; Mukherjee, J.; Wlodkowski, A.; Karunakaran, E.; Biggs, C.A.; Wright, P.C.; Albers, S.-V. Macromolecular fingerprinting of sulfolobus species in biofilm: A transcriptomic and proteomic approach combined with spectroscopic analysis. J. Proteome Res. 2011, 10, 4105–4119. [Google Scholar] [CrossRef] [PubMed]
  330. Koerdt, A.; Gödeke, J.; Berger, J.; Thormann, K.M.; Albers, S.-V. Crenarchaeal biofilm formation under extreme conditions. PLoS ONE 2010, 5. [Google Scholar] [CrossRef] [PubMed]
  331. Pohlschroder, M.; Esquivel, R.N. Archaeal type IV pili and their involvement in biofilm formation. Front. Microbiol. 2015, 6, 190. [Google Scholar] [CrossRef] [PubMed]
  332. Orell, A.; Peeters, E.; Vassen, V.; Jachlewski, S.; Schalles, S.; Siebers, B.; Albers, S.-V. Lrs14 transcriptional regulators influence biofilm formation and cell motility of Crenarchaea. ISME J. 2013, 7, 1886–1898. [Google Scholar] [CrossRef] [PubMed]
  333. Baker-Austin, C.; Potrykus, J.; Wexler, M.; Bond, P.L.; Dopson, M. Biofilm development in the extremely acidophilic archaeon “Ferroplasma acidarmanus” Fer1. Extremophiles 2010, 14, 485–491. [Google Scholar] [CrossRef] [PubMed]
  334. Zhang, R.Y.; Neu, T.R.; Bellenberg, S.; Kuhlicke, U.; Sand, W.; Vera, M. Use of lectins to in situ visualize glycoconjugates of extracellular polymeric substances in acidophilic archaeal biofilms. Microb. Biotechnol. 2015, 8, 448–461. [Google Scholar] [CrossRef] [PubMed]
  335. Poli, A.; di Donato, P.; Abbamondi, G.R.; Nicolaus, B. Synthesis, production, and biotechnological applications of exopolysaccharides and polyhydroxyalkanoates by Archaea. Archaea 2011, 2011, 693253. [Google Scholar] [CrossRef] [PubMed]
  336. Fröls, S. Archaeal biofilms: Widespread and complex. Biochem. Soc. Trans. 2013, 41, 393–398. [Google Scholar] [CrossRef] [PubMed]
  337. Jarrell, K.; Ding, Y.; Nair, D.; Siu, S. Surface Appendages of Archaea: Structure, Function, Genetics and Assembly. Life 2013, 3, 86–117. [Google Scholar] [CrossRef] [PubMed]
  338. Orell, A.; Fröls, S.; Albers, S.-V. Archaeal biofilms: The great unexplored. Annu. Rev. Microbiol. 2013, 67, 337–354. [Google Scholar] [CrossRef] [PubMed]
  339. Miot, J.; Benzerara, K.; Kappler, A. Investigating Microbe-Mineral Interactions: Recent Advances in X-Ray and Electron Microscopy and Redox-Sensitive Methods. Annu. Rev. Earth Planet. Sci. 2014, 42, 271–289. [Google Scholar] [CrossRef]
  340. Gralnick, J.A.; Newman, D.K. Extracellular respiration. Mol. Microbiol. 2007, 65, 1–11. [Google Scholar] [CrossRef] [PubMed]
  341. Gray, H.B.; Winkler, J.R. Electron tunneling through proteins. Q. Rev. Biophys. 2003, 36, 341–372. [Google Scholar] [CrossRef] [PubMed]
  342. Richardson, D.J.; Butt, J.N.; Fredrickson, J.K.; Zachara, J.M.; Shi, L.; Edwards, M.J.; White, G.; Baiden, N.; Gates, A.J.; Marritt, S.J.; et al. The “porin-cytochrome” model for microbe-to-mineral electron transfer. Mol. Microbiol. 2012, 85, 201–212. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  343. White, G.F.; Shi, Z.; Shi, L.; Wang, Z.; Dohnalkova, A.C.; Marshall, M.J.; Fredrickson, J.K.; Zachara, J.M.; Butt, J.N.; Richardson, D.J.; et al. A Rapid electron exchange between surface-exposed bacterial cytochromes and Fe(III) minerals. Proc. Natl. Acad. Sci. USA 2013, 110, 6346–6351. [Google Scholar] [CrossRef] [PubMed]
  344. El-Naggar, M.; Finkel, S. Live Wires. Scientist 2013, 27, 38–43. [Google Scholar]
  345. Marsili, E.; Baron, D.B.; Shikhare, I.D.; Coursolle, D.; Gralnick, J.A.; Bond, D.R. Shewanella secretes flavins that mediate extracellular electron transfer. Proc. Natl. Acad. Sci. USA 2008, 105, 3968–3973. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  346. Von Canstein, H.; Ogawa, J.; Shimizu, S.; Lloyd, J.R. Secretion of flavins by Shewanella species and their role in extracellular electron transfer. Appl. Environ. Microbiol. 2008, 74, 615–623. [Google Scholar] [CrossRef] [PubMed]
  347. Brutinel, E.D.; Gralnick, J.A. Shuttling happens: Soluble flavin mediators of extracellular electron transfer in Shewanella. Appl. Microbiol. Biotechnol. 2012, 93, 41–48. [Google Scholar] [CrossRef] [PubMed]
  348. Reguera, G.; McCarthy, K.D.; Mehta, T.; Nicoll, J.S.; Tuominen, M.T.; Lovley, D.R. Extracellular electron transfer via microbial nanowires. Nature 2005, 435, 1098–1101. [Google Scholar] [CrossRef] [PubMed]
  349. Gorby, Y.A.; Yanina, S.; McLean, J.S.; Rosso, K.M.; Moyles, D.; Dohnalkova, A.; Beveridge, T.J.; Chang, I.S.; Kim, B.H.; Kim, K.S.; et al. Electrically conductive bacterial nanowires produced by Shewanella oneidensis strain MR-1 and other microorganisms. Proc. Natl. Acad. Sci. USA 2006, 103, 11358–11363. [Google Scholar] [CrossRef] [PubMed]
  350. El-Naggar, M.Y.; Wanger, G.; Leung, K.M.; Yuzvinsky, T.D.; Southam, G.; Yang, J.; Lau, W.M.; Nealson, K.H.; Gorby, Y.A. Electrical transport along bacterial nanowires from Shewanella oneidensis MR-1. Proc. Natl. Acad. Sci. USA 2010, 107, 18127–18131. [Google Scholar] [CrossRef] [PubMed]
  351. Lovley, D.R. Long-range electron transport to Fe(III) oxide via pili with metallic-like conductivity. Biochem. Soc. Trans. 2012, 40, 1186–1190. [Google Scholar] [CrossRef] [PubMed]
  352. Pirbadian, S.; Barchinger, S.E.; Leung, K.M.; Byun, H.S.; Jangir, Y.; Bouhenni, R.A.; Reed, S.B.; Romine, M.F.; Saffarini, D.A.; Shi, L.; et al. Shewanella oneidensis MR-1 nanowires are outer membrane and periplasmic extensions of the extracellular electron transport components. Proc. Natl. Acad. Sci. USA 2014, 111, 1–6. [Google Scholar] [CrossRef] [PubMed]
  353. Gorby, Y.; Mclean, J.; Korenevsky, A.; Rosso, K.; El-Naggar, M.Y.; Beveridge, T.J. Redox-reactive membrane vesicles produced by Shewanella. Geobiology 2008, 6, 232–241. [Google Scholar] [CrossRef] [PubMed]
  354. Snider, R.M.; Strycharz-Glaven, S.M.; Tsoi, S.D.; Erickson, J.S.; Tender, L.M. Long-range electron transport in Geobacter sulfurreducens biofilms is redox gradient-driven. Proc. Natl. Acad. Sci. USA 2012, 109, 15467–15472. [Google Scholar] [CrossRef] [PubMed]
  355. Pfeffer, C.; Larsen, S.; Song, J.; Dong, M.; Besenbacher, F.; Meyer, R.L.; Kjeldsen, K.U.; Schreiber, L.; Gorby, Y.A.; El-Naggar, M.Y.; et al. Filamentous bacteria transport electrons over centimetre distances. Nature 2012, 491, 218–221. [Google Scholar] [CrossRef] [PubMed]
  356. Malvankar, N.S.; Vargas, M.; Nevin, K.P.; Franks, A.E.; Leang, C.; Kim, B.; Inoue, K.; Mester, T.; Covalla, S.F.; Johnson, J.P.; et al. Tunable metallic-like conductivity in microbial nanowire networks. Nat. Nanotechnol. 2011, 6, 573–579. [Google Scholar] [CrossRef] [PubMed]
  357. Vargas, M.; Malvankar, N.S.; Tremblay, P.L.; Leang, C.; Smith, J.A.; Patel, P.; Synoeyenbos-West, O.; Nevin, K.P.; Lovley, D.R. Aromatic amino acids required for pili conductivity and long-range extracellular electron transport in Geobacter sulfurreducens. MBio 2013, 4. [Google Scholar] [CrossRef]
  358. Pirbadian, S.; El-Naggar, M.Y. Multistep hopping and extracellular charge transfer in microbial redox chains. Phys. Chem. Chem. Phys. 2012, 14, 13802. [Google Scholar] [CrossRef] [PubMed]
  359. Polizzi, N.F.; Skourtis, S.S.; Beratan, D.N. Physical constraints on charge transport through bacterial nanowires. Faraday Discuss. 2012, 155, 43. [Google Scholar] [CrossRef] [PubMed]
  360. Strycharz-Glaven, S.M.; Tender, L.M. Reply to the “Comment on ‘On electrical conductivity of microbial nanowires and biofilms’” N.S. Malvankar, M.T. Tuominen and D.R. Lovley. Energy Environ. Sci., 2012, 5, doi:10.1039/c2ee02613a. Energy Environ. Sci. 2012, 5, 6250–6255. [Google Scholar] [CrossRef]
  361. Li, Y.; Li, H. Type IV pili of Acidithiobacillus ferrooxidans can transfer electrons from extracellular electron donors. J. Basic Microbiol. 2014, 54, 226–231. [Google Scholar] [CrossRef] [PubMed]
  362. Rojas-Chapana, J.A.; Tributsch, H. Interfacial activity and leaching patterns of Leptospirillum ferrooxidans on pyrite. FEMS Microbiol. Ecol. 2004, 47, 19–29. [Google Scholar] [CrossRef]
  363. Medvedev, D.; Stuchebrukhov, A.A. DNA repair mechanism by photolyase: Electron transfer path from the photolyase catalytic cofactor FADH(-) to DNA thymine dimer. J. Theor. Biol. 2001, 210, 237–248. [Google Scholar] [CrossRef] [PubMed]
  364. Taylor, E.S.; Lower, S.K. Thickness and surface density of extracellular polymers on Acidithiobacillus ferrooxidans. Appl. Environ. Microbiol. 2008, 74, 309–311. [Google Scholar] [CrossRef] [PubMed]
  365. Rojas-Chapana, J.A.; Tributsch, H. Bio-leaching of pyrite accelerated by cysteine. Process Biochem. 2000, 35, 815–824. [Google Scholar] [CrossRef]
  366. Rojas-Chapana, J.A.; Tributsch, H. Biochemistry of sulfur extraction in bio-corrosion of pyrite by Thiobacillus ferrooxidans. Hydrometallurgy 2001, 59, 291–300. [Google Scholar] [CrossRef]
  367. Blake, R.C.; Sasaki, K.; Ohmura, N. Does aporusticyanin mediate the adhesion of Thiobacillus ferrooxidans to pyrite? Hydrometallurgy 2001, 59, 357–372. [Google Scholar] [CrossRef]
  368. Kuehn, M.J.; Kesty, N.C. Bacterial outer membrane vesicles and the host—Pathogen interaction. Genes Dev. 2005, 19, 2645–2655. [Google Scholar] [CrossRef] [PubMed]
  369. Mashburn-Warren, L.; Mclean, R.J.C.; Whiteley, M. Gram-negative outer membrane vesicles: Beyond the cell surface. Geobiology 2008, 6, 214–219. [Google Scholar] [CrossRef] [PubMed]
  370. Ellen, A.F.; Albers, S.V.; Huibers, W.; Pitcher, A.; Hobel, C.F.V.; Schwarz, H.; Folea, M.; Schouten, S.; Boekema, E.J.; Poolman, B.; et al. Proteomic analysis of secreted membrane vesicles of archaeal Sulfolobus species reveals the presence of endosome sorting complex components. Extremophiles 2009, 13, 67–79. [Google Scholar] [CrossRef] [PubMed]
  371. Prangishvili, D.; Holz, I.; Stieger, E.; Nickell, S.; Kristjansson, J.K.; Zillig, W. Sulfolobicins, specific proteinaceous toxins produced by strains of the extremely thermophilic archaeal genus Sulfolobus. J. Bacteriol. 2000, 182, 2985–2988. [Google Scholar] [CrossRef] [PubMed]
  372. Ellen, A.F.; Zolghadr, B.; Driessen, A.M.J.; Albers, S.-V. Shaping the archaeal cell envelope. Archaea 2010, 2010, 608243. [Google Scholar] [CrossRef] [PubMed]
  373. Ellen, A.F.; Albers, S.-V.; Driessen, A.J.M. Comparative study of the extracellular proteome of Sulfolobus species reveals limited secretion. Extremophiles 2010, 14, 87–98. [Google Scholar] [CrossRef] [PubMed]
  374. Roden, E.E. Microbiological controls on geochemical kinetics 2: Case study on microbial oxidation of metal sulfide minerals and future prospects. In Kinetics of Water-Rock Interaction; Brantley, S.L., Kubicki, J.D., White, A.F., Eds.; Springer: New York, NY, USA, 2008; pp. 417–467. [Google Scholar]
  375. Johnson, D.B. Biomining-biotechnologies for extracting and recovering metals from ores and waste materials. Curr. Opin. Biotechnol. 2014, 30, 24–31. [Google Scholar] [CrossRef] [PubMed]
  376. Watling, H.R. The bioleaching of sulphide minerals with emphasis on copper sulphides—A review. Hydrometallurgy 2006, 84, 81–108. [Google Scholar] [CrossRef]
  377. Watling, H.R. Review of Biohydrometallurgical Metals Extraction from Polymetallic Mineral Resources. Minerals 2014, 5, 1–60. [Google Scholar] [CrossRef]
  378. Du Plessis, C.A.; Batty, J.D.; Dew, D.W. Commercial Applications of Thermophile Bioleaching. In Biomining; Rawlings, D.E., Johnson, D.B., Eds.; Springer-Verlag: Berlin, Germany, 2007; pp. 57–80. [Google Scholar]
  379. Neale, J.W.; Robertson, S.W.; Muller, H.H.; Gericke, M. Integrated piloting of a thermophilic bioleaching process for the treatment of a low-grade nickel-copper sulphide concentrate. J. South. African Inst. Min. Metall. 2009, 109, 273–293. [Google Scholar]
  380. Batty, J.D.; Rorke, G.V. Development and commercial demonstration of the BioCOP™ thermophile process. Hydrometallurgy 2006, 83, 83–89. [Google Scholar] [CrossRef]
  381. Logan, T.C.; Seal, T.; Brierley, J.A. Whole-ore heap biooxidation of sulfidic gold-bearing ores. In Biomining; Rawlings, D.E., Johnson, D.B., Eds.; Springer-Verlag: Berlin, Germany, 2007; pp. 113–138. [Google Scholar]
  382. Dew, D.W.; Buuren, C.; van Mcewan, K.; Bowker, C. Bioleaching of base metal sulphide concentrates: A comparison of high and low temperature bioleaching. J. S. Afr. Inst. Min. Metall. 2000, 100, 409–414. [Google Scholar]
  383. Khoshkhoo, M.; Dopson, M.; Shchukarev, A.; Sandström, Å. Chalcopyrite leaching and bioleaching: An X-ray photoelectron spectroscopic (XPS) investigation on the nature of hindered dissolution. Hydrometallurgy 2014, 149, 220–227. [Google Scholar] [CrossRef]
  384. Gericke, M.; Govender, Y.; Pinches, A. Tank bioleaching of low-grade chalcopyrite concentrates using redox control. Hydrometallurgy 2010, 104, 414–419. [Google Scholar] [CrossRef]
  385. Ahonen, L.; Tuovinen, O.H. Silver catalysis of the bacterial leaching of chalcopyrite-containing ore material in column reactors. Miner. Eng. 1990, 3, 437–445. [Google Scholar] [CrossRef]
  386. Crundwell, F.K. The semiconductor mechanism of dissolution and the pseudo-passivation of chalcopyrite. Can. Metall. Q. 2015. [Google Scholar] [CrossRef]
  387. Abdollahi, H.; Shafaei, S.Z.; Noaparast, M.; Manafi, Z.; Niemelä, S.I.; Tuovinen, O.H. Mesophilic and thermophilic bioleaching of copper from a chalcopyrite-containing molybdenite concentrate. Int. J. Miner. Process. 2014, 128, 25–32. [Google Scholar] [CrossRef]
  388. Romano, P.; Blázquez, M.L.; Alguacil, F.J.; Muñoz, J.A.; Ballester, A.; González, F. Comparative study on the selective chalcopyrite bioleaching of a molybdenite concentrate with mesophilic and thermophilic bacteria. FEMS Microbiol. Lett. 2001, 196, 71–75. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  389. Takatsugi, K.; Sasaki, K.; Hirajima, T. Mechanism of the enhancement of bioleaching of copper from enargite by thermophilic iron-oxidizing archaea with the concomitant precipitation of arsenic. Hydrometallurgy 2011, 109, 90–96. [Google Scholar] [CrossRef]
  390. D’Hughes, P.D.; Foucher, S. HIOX® Project: A bioleaching process for the treatment of chalcopyrite concentrates using extreme thermophilesthermophiles. In Biohydrometallurgy: Fundamentals, Technology and Sustainable Development; Ciminelli, V.S.T., Garcia, O., Eds.; Elsevier: Amsterdam, The Netherlands, 2001; pp. 75–83. [Google Scholar]
  391. Marsh, R.M.; Norris, P.R. The isolation of some thermophilic, autotrophic, iron- and sulphur-oxidizing bacteria. FEMS Microbiol. Lett. 1983, 17, 311–315. [Google Scholar] [CrossRef]
  392. Le Roux, N.W.; Wakerley, W. Leaching of Chalcopyrite (CuFeS2) at 70 °C using Sulfolobus. In Proceedings of Biohydrometallurgy 87’; Norris, P.R., Kelly, D.P., Eds.; Science and Technology Letters: Surrey, UK, 1998; pp. 305–317. [Google Scholar]
  393. Lindstrom, E.B.; Gunneriusson, L. Thermophilic bioleaching of arsenopyrite using Sulfolobus and a semi-continuous laboratory procedure. J. Ind. Microbiol. 1990, 5, 375–382. [Google Scholar] [CrossRef]
  394. Gericke, M.; Pinches, A.; van Rooyen, J. Bioleaching of a chalcopyrite concentrate using an extremely thermophilic culture. Int. J. Miner. Process. 2001, 62, 243–255. [Google Scholar] [CrossRef]
  395. Konishi, Y.; Yoshida, S.; Asai, S. Bioleaching of pyrite by acidophilic thermophile Acidianus brierleyi. Biotechnol. Bioeng. 1995, 48, 592–600. [Google Scholar] [CrossRef] [PubMed]
  396. Sandström, Å.; Petersson, S. Bioleaching of a complex sulphide ore with moderate thermophilic and extreme thermophilic microorganisms. Hydrometallurgy 1997, 46, 181–190. [Google Scholar]
  397. Harvey, T.J.; Holder, N.; Stanek, T. Thermophilic Bioheap Leaching of Chalcopyrite Concentrates. Eur. J. Miner. Process. Environ. Prot. 2002, 2, 253–263. [Google Scholar]
  398. Hita, R.; Wang, H.; Bigham, J.M.; Torrent, J.; Tuovinen, O.H. Bioleaching of a pyritic sludge from the Aznalcóllar (Spain) mine spillage at ambient and elevated temperatures. Hydrometallurgy 2008, 93, 76–79. [Google Scholar] [CrossRef]
  399. Bharadwaj, A.; Ting, Y.P. Bioleaching of spent hydrotreating catalyst by acidophilic thermophile Acidianus brierleyi: Leaching mechanism and effect of decoking. Bioresour. Technol. 2013, 130, 673–680. [Google Scholar] [CrossRef] [PubMed]
  400. Vukovic, M.; Strbac, N.; Sokic, M.; Grekulovic, V.; Cvetkovski, V. Bioleaching of pollymetallic sulphide concentrate using thermophilic bacteria. Hem. Ind. 2014, 68, 575–583. [Google Scholar] [CrossRef] [Green Version]
  401. Clark, M.E.; Batty, J.D.; van Buuren, C.B.; Dew, D.W.; Eamon, M.A. Biotechnology in minerals processing: Technological breakthroughs creating value. Hydrometallurgy 2006, 83, 3–9. [Google Scholar] [CrossRef]
  402. Van Staden, P.J.; Gericke, M.; Craven, P.M. Minerals biotechnology: Trends, opportunities and challenges. In Hydrometallurgy 2008 : Proceedings of the Sixth International Symposium; Society for Mining Metallurgy and Exploration: Littleton, CO, USA, 2008; pp. 6–13. [Google Scholar]
  403. Petersen, J.; Dixon, D.G. Competitive bioleaching of pyrite and chalcopyrite. Hydrometallurgy 2006, 83, 40–49. [Google Scholar] [CrossRef]
  404. Javadi Nooshabadi, A.; Hanumantha Rao, K. Formation of hydrogen peroxide by sulphide minerals. Hydrometallurgy 2014, 141, 82–88. [Google Scholar] [CrossRef]
  405. Jones, G.C.; van Hille, R.P.; Harrison, S.T.L. Reactive oxygen species generated in the presence of fine pyrite particles and its implication in thermophilic mineral bioleaching. Appl. Microbiol. Biotechnol. 2013, 97, 2735–2742. [Google Scholar] [CrossRef] [PubMed]
  406. Langwaldt, J. Bioleaching of multimetal black shale by thermophilic micro-organisms. Adv. Mater. Res. 2007, 20, 167. [Google Scholar] [CrossRef]
  407. Mishra, D.; Rhee, Y.H. Microbial leaching of metals from solid industrial wastes. J. Microbiol. 2014, 52, 1–7. [Google Scholar] [CrossRef] [PubMed]
  408. Lee, J.C.; Pandey, B.D. Bio-processing of solid wastes and secondary resources for metal extraction—A review. Waste Manag. 2012, 32, 3–18. [Google Scholar] [CrossRef] [PubMed]
  409. Reed, C. Earth microbe prefers living on meteorites. Science 2015. [Google Scholar] [CrossRef]
  410. Takahashi, Y.; Châtellier, X.; Hattori, K.H.; Kato, K.; Fortin, D. Adsorption of rare earth elements onto bacterial cell walls and its implication for REE sorption onto natural microbial mats. Chem. Geol. 2005, 219, 53–67. [Google Scholar] [CrossRef]
  411. Tsuruta, T. Accumulation of Rare Earth Elements in Various Microorganisms. J. Rare Earths 2007, 25, 526–532. [Google Scholar] [CrossRef]
  412. Emmanuel, E.S.C.; Ananthi, T.; Anandkumar, B.; Maruthamuthu, S. Accumulation of rare earth elements by siderophore-forming Arthrobacter luteolus isolated from rare earth environment of Chavara, India. J. Biosci. 2012, 37, 25–31. [Google Scholar] [CrossRef] [PubMed]
  413. Horiike, T.; Yamashita, M. A new fungal isolate, Penidiella sp. Strain T9, accumulates the rare earth element dysprosium. Appl. Environ. Microbiol. 2015, 81, 3062–3068. [Google Scholar] [CrossRef] [PubMed]
  414. Pol, A.; Barends, T.R.M.; Dietl, A.; Khadem, A.F.; Eygensteyn, J.; Jetten, M.S.M.; Op den Camp, H.J.M. Rare earth metals are essential for methanotrophic life in volcanic mudpots. Environ. Microbiol. 2014, 16, 255–264. [Google Scholar] [CrossRef] [PubMed]
  415. Cvetkovic, A.; Menon, A.L.; Thorgersen, M.P.; Scott, J.W.; Poole, F.L.; Jenney, F.E.; Lancaster, W.A.; Praissman, J.L.; Shanmukh, S.; Vaccaro, B.J.; et al. Microbial metalloproteomes are largely uncharacterized. Nature 2010, 466, 779–782. [Google Scholar] [CrossRef] [PubMed]

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MDPI and ACS Style

Wheaton, G.; Counts, J.; Mukherjee, A.; Kruh, J.; Kelly, R. The Confluence of Heavy Metal Biooxidation and Heavy Metal Resistance: Implications for Bioleaching by Extreme Thermoacidophiles. Minerals 2015, 5, 397-451.

AMA Style

Wheaton G, Counts J, Mukherjee A, Kruh J, Kelly R. The Confluence of Heavy Metal Biooxidation and Heavy Metal Resistance: Implications for Bioleaching by Extreme Thermoacidophiles. Minerals. 2015; 5(3):397-451.

Chicago/Turabian Style

Wheaton, Garrett, James Counts, Arpan Mukherjee, Jessica Kruh, and Robert Kelly. 2015. "The Confluence of Heavy Metal Biooxidation and Heavy Metal Resistance: Implications for Bioleaching by Extreme Thermoacidophiles" Minerals 5, no. 3: 397-451.

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