Carbon disulfide. Just toxic or also bioregulatory and/or therapeutic?

Abstract

The overview presented here has the goal of examining whether carbon disulfide (CS₂) may play a role as an endogenously generated bioregulator and/or has therapeutic value. The neuro- and reproductive system toxicity of CS₂ has been documented from its long-term use in the viscose rayon industry. CS₂ is also used in the production of dithiocarbamates (DTCs), which are potent fungicides and pesticides, thus raising concern that CS₂ may be an environmental toxin. However, DTCs also have recognized medicinal use in the treatment of heavy metal poisonings as well as having potency for reducing inflammation. Three known small molecule bioregulators (SMBs) nitric oxide, carbon monoxide, and hydrogen sulfide were initially viewed as environmental toxins. Yet each is now recognized as having intricate, though not fully elucidated, biological functions at concentration regimes far lower than the toxic doses. The literature also implies that the mammalian chemical biology of CS₂ has broader implications from inflammatory states to the gut microbiome. On these bases, we suggest that the very nature of CS₂ poisoning may be related to interrupting or overwhelming relevant regulatory or signaling process(es), much like other SMBs.

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Anthony W. DeMartino

Anthony (Tony) DeMartino received his BS in biochemistry from Wake Forest University in Winston-Salem, NC where he worked with Prof. Ulrich Bierbach. He is currently finishing his PhD in chemistry with Prof. Peter C. Ford at UC Santa Barbara where he has worked on photochemical NO donors and is currently examining the potential of CS2 as a small molecule bioregulator through the development of effective donors and various collaborative pharmacological studies.

David F. Zigler

David F. Zigler is an Assistant Professor in the Department of Chemistry & Biochemistry at California Polytechnic State University in San Luis Obispo, CA. After his graduate work with the late Karen J. Brewer at Virginia Tech, he worked as a postdoc in the Ford lab at UC Santa Barbara to create new compounds with caged CS2. Appointments at the University of North Carolina followed with Malcolm Forbes using EPR to study reactions with photo-generated ^1O2 and with John Papanikolas in the UNC-EFRC Center for Solar Fuels to examine photo-initiated electron transfer at semiconductor/electrolyte interfaces. Professor Zigler's current research interests include inorganic phototherapeutics and materials for solar-to-chemical energy conversion.

Jon M. Fukuto

Jon M. Fukuto received a BA in chemistry from UC San Diego and PhD in chemistry from UC Berkeley where he worked with Prof. Fritz Jensen. He then did postdoctoral work at UCLA in Pharmacology with Prof. Arthur Cho. After a short stint in industry, he returned to UCLA as faculty in Pharmacology where he resided for over 18 years. At UCLA he worked on the chemical biology of the small molecule signaling agents nitric oxide (NO), nitroxyl (HNO) and hydrogen sulfide (H2S). He is currently a Professor of Chemistry at Sonoma State University where he continues this work.

Peter C. Ford

Professor Peter C. Ford joined the UCSB faculty after graduate study with Ken Wiberg at Yale and a postdoctoral fellowship with Henry Taube at Stanford. He has been a Visiting Fellow at the Australian National U, Guest Professor at U Copenhagen, Humboldt Fellow at U Regensberg and U Muenster and Guest Investigator at the US National Cancer Institute. Honors include a Humboldt Senior US Scientist Award, Inter-American Photochemical Society 2008 Award in Photochemistry, ACS 2013 National Award for Distinguished Service to Inorganic Chemistry and RSC 2015 Award in Inorganic Mechanisms. He is an AAAS Fellow and a RSC Fellow. His current research focuses on photochemical delivery of bioactive small molecules to physiological targets and biomass conversion to chemicals and fuels. He has served as research advisor to 65 PhD graduates and a number of postdocs, undergraduates and international visiting scholars.

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1. Introduction

Small molecule bioregulators (SMBs) are physiological signaling agents with interconnected pathways based on their unique chemistry and interactions with specific biological targets.¹ This cellular signaling paradigm is a burgeoning field in mammalian chemical biology, and over the past several decades, SMBs central to this network have been identified. Three key SMBs, nitric oxide (NO), carbon monoxide (CO) and hydrogen sulfide (H₂S) have been called “gasotransmitters” although they are solubilized at the physiologically pertinent concentrations. A number of specific functions have been elucidated; for example, NO plays a key role in regulating blood pressure via the activation of soluble-guanylyl cyclase (sGC).² Other known physiological functions are not yet as well understood, and the biological roles of these SMBs and of respective derivatives such as nitroxyl (HNO), peroxynitrite (ONOO⁻) and persulfides (R–SS⁻) continue to be the subjects of extensive biomedical research efforts.^1–11

If one were to envision an unidentified (so far) SMB in mammalian chemical biology, it may be appropriate to consider the shared properties of NO, CO, and H₂S. All are relatively nonpolar but are partially soluble in aqueous and lipid systems; thus, they readily diffuse in physiological media via spatial concentration gradients.¹² The actions of these SMBs involve chemical reactions with targets such as metal centers or redox active amino acids.⁷ Furthermore, all were initially considered to be environmental toxins; yet each was found to be formed by endogenous processes. In this context, carbon disulfide (CS₂) is a small, nonpolar, readily diffusible molecule long recognized for its toxicity that may be generated endogenously. These similarities led us to contemplate whether CS₂ might be an unrecognized SMB and/or have therapeutic potential.¹³

The purpose of this review is to explore the possibility of such roles for carbon disulfide. We will first outline possible modes of toxicity and then will survey the chemical properties of CS₂ in the context of physiologically relevant conditions. Next, we will discuss known mechanisms for CS₂ metabolism in order to visualize a framework by which CS₂ may be managed endogenously. Finally, some therapeutic targets are suggested.

2. General physical and chemical properties of CS₂

2.1. Physical properties and known sources

Pure CS₂ is a colorless, pleasant-smelling liquid displaying a high vapor pressure (48.21 kPa at 25 °C), moderate water solubility (2.3 g L⁻¹ at 22 °C), and high lipophilicity.¹⁴ It is a linear, nonpolar molecule like its homolog carbon dioxide (CO₂) with carbon–sulfur bonds 155.6 pm in length.¹⁵ CS₂ is slightly more soluble than CO₂ in aqueous solutions; the Henry's law constants at 25 °C in pure water are 0.054 M atm⁻¹ and 0.034 M atm⁻¹ respectively.¹⁶

Although CS₂ is a listed environmental pollutant, it is produced naturally by soil-based microorganisms and is formed in vegetation fires and volcanoes. These sources account for 40–50% of atmospheric release, and the rest is anthropogenic.¹⁷ CS₂ is produced industrially from heating a carbon source such as charcoal or natural gas with sulfur.¹⁸ The annual industrial production is ∼75 million kilograms per year (ca. 2010) for use in the rayon and cellophane, oil and gas, agricultural, and metal ore processing industries.^18,19 It has been used in production of viscose rayon since the nineteenth century,²⁰ when the deleterious effects on exposed workers in that industry were first described.²¹

2.2. Toxicity

As noted above, CS₂ toxicity was first described in the 1850s²¹ and is the subject of numerous subsequent studies.^22–26 Of particular interest are the neurotoxic effects.^27–33 Acute CS₂ poisoning causes toxic psychosis and narcotic somnolence. Long-term sub-acute exposures are linked with memory, behavioral and cognitive disturbances, and also concomitant nerve demyelination, cytostructural damage, and progressive paralysis of the extremities.^33,34 The mechanism of CS₂ neurotoxicity, however, is currently unknown. Epidemiological studies of chronic CS₂ exposure are somewhat contradictory, particularly regarding reproductive^35–37 and cardiovascular issues.^38,39 From a brief overview of the literature, the described toxicity, pathology, and/or metabolism of CS₂ indicate a breadth of targeted biochemical effects.

2.3. Some key chemical reactions

CS₂ is stable to hydration and at biologically relevant pH values is relatively unreactive toward hydrolysis to carbonyl sulfide (COS) in aqueous solutions (eqn (1)).The approximate half-life of this reaction is 1.1 years in pH 9 aqueous solution.^19,40 CS₂ does react with a variety of stronger nucleophiles. For example in strongly alkaline solution, it forms trithiocarbonate and carbonate (eqn (2)).⁴¹

The industrially important reaction of CS₂ with primary and secondary alcohols and a metal hydroxide to form xanthates (eqn (3)) was reported as early as 1822.⁴²

Acid catalyzes xanthate decomposition to the alcohol and CS₂, while in neutral or slightly alkaline pH xanthates undergo hydrolytic disproportionation (eqn (4)).⁴¹ Hydrolytic disproportionation is slow with half-lives on the order of several hours.⁴³

The analogous reactions of CS₂ with primary and secondary amines lead to the formation of dithiocarbamates (DTCs) (eqn (5)). This equilibrium is dependent on the nature of the amine (e.g., sterics and electronics) and on the solution pH. In acidic solutions, dithiocarbamates readily decompose to the constituent amine plus CS₂.^44–46 At pH 7.4 and 35 °C, primary and most secondary alkyl-DTCs are relatively stable, whereas under mild acidic conditions, secondary alkyl-DTCs revert to the amine and CS₂ with rate constants on the order of 10⁻³ s⁻¹ at pH 5.9.⁴⁷

A CS₂ reaction that might have particular significance with regard to biological properties is the reversible formation of trithiocarbonates (TTCs, also known as thioxanthates) as illustrated in eqn (6).^48–50 We will illustrate possible roles of this reaction in greater detail below.

2.4. Interactions with metal centers

Carbon disulfide and its adducts (xanthates, DTCs, and TTCs) can serve as ligands in metal complexes. Typically, CS₂ coordinates more strongly to metal centers than does either CO₂ or COS.^51,52 There are a number of examples of η¹- and η²-CS₂ complexes^53–55 as well as polynuclear complexes⁵⁶ where the CS₂ moiety is a bridging ligand (Fig. 1).

Fig. 1 Binding modes of CS₂ with metal centers.

Such coordination can activate CS₂ toward reaction with nucleophiles. For example, reversible CS₂ insertion into a metal–ligand bond is a common reaction (eqn (7)), presumably because of the stronger bonding of the 1,1-dithiolate (–CS₂⁻) functionality.

This has been seen for metal-bound thiolates⁵⁷ and alkoxides⁵⁸ and with thiols or amines with loss of H⁺.⁵⁹ Analogous reactions of CS₂ with coordinated nucleophilic carbon centers result in C–C bond formation via the generation of a dithiocarboxylate ligand.⁶⁰ The analogous reactions with coordinated phosphines lead to a zwitterion (R₃P⁺–CS₂⁻) adducts as a bidentate ligand bound to the metal.⁶¹ Similar CS₂ insertions have been reported for Cu(I)–amidinate complexes⁶² and for nickel(II)–amine bonds in polar protic or aprotic solvents (eqn (8)).⁶³ For example, although the trithiocarbonate zwitterion [⁺H₃NCH₂CH₂SCS₂⁻] forms in the direct CS₂ reaction with 2-aminoethanethiol,⁴⁸ the reaction with bis(2-mercapto-ethylamine)nickel(II) favors insertion between the metal and the amine to give a pendant thiol on the DTC complex.⁶³

Metal centers also react with various RXCS₂⁻ (where X = NR′ or S) ligands to form stable complexes under a variety of conditions. Although generally stable in neutral aqueous solutions, some M(S₂CXR) complexes eliminate CS₂.^64–69 For example, spontaneous release of CS₂ from the Ni(II) S-benzyl- and tert-butyl-trithiocarbonate complexes (eqn (9)) occurs with first order rate constants on the order of 10⁻⁴ s⁻¹ at 35 °C in lipophilic media.⁶⁴

Studies of such ligand–metal interactions involving 1,1-dithiolate functional groups (–CS₂⁻) have received relatively low attention in recent years but have been the subject of several reviews.^70–73

3. Biological chemistry

3.1. Hydrolysis (or not)

Like CO₂, carbon disulfide is rather unreactive toward spontaneous hydration and hydrolysis at biological pH values, so a significant proportion of the dissolved compound is present in the solvated form, CS₂ (aq). Furthermore, carbonic anhydrase (CA), the enzyme that catalyzes the hydration of CO₂, is ineffective in activating the analogous reaction of CS₂ (eqn (1)). For example, with excess CS₂, the turnover frequency of cytosolic CA II, the most efficient of the fourteen CA isomers,⁷⁴ is about 9 orders of magnitude slower (∼5 × 10⁻⁴ s⁻¹) than the analogous reaction with CO₂ (∼10⁶ s⁻¹).⁷⁵

The first step of CS₂ hydrolysis, like CO₂, is hydration. Using a model for the Zn-active site of CA II, Anders and coworkers computationally probed the formation of dithiocarbonate, the hydrated intermediate of CS₂ (Scheme 1) by considering a mechanism analogous to CO₂ hydration.^76–78 It is argued that dithiocarbonate formation has a slightly higher barrier of activation owing to a decreased electrophilicity of CS₂.⁷⁶ In addition, the intermediate dithiobicarbonate (HCS₂O⁻) is a thermodynamic sink, which further slows hydrolysis.⁵¹ The calculations with this active site model suggest that CA does not complete a catalytic cycle with CS₂. Rather, CS₂ is proposed to inhibit CA. Although CS₂ does not appear to interact with CA II,⁷⁵ xanthates and TTCs are effective inhibitors.⁷⁹ Nevertheless, there are bacterial-evolved, archaeal CS₂ hydrolases that do convert CS₂ to CO₂ and H₂S through intermediate COS (eqn (1) and (10)).⁸⁰ In spite of the high homology with carbonic anhydrases, these CS₂ hydrolases apparently do not use CO₂ as a substrate,⁸⁰ just as CA does not efficiently catalyze CS₂ hydrolysis. Notably, COS is readily hydrolyzed by both types.^75,77,78

Scheme 1 Hydration of CS₂ in Zn-based CA model active site. Anders and coworkers calculated a mechanism analogous to that for CO₂, with hydration leading to a stable dithiobicarbonate adduct. The subsequent step (hydrolysis) is slow.

The presence of CS₂ hydrolases in some microorganisms and clear divergence in evolution imply an evolutionary need for a CS₂-specific enzyme. Thiobacillus thioparus and Thiobacillus denitrificans are microbes that use reduced sulfur species (dimethyl sulfide, methyl mercaptan, and H₂S) for energy production.⁸¹ These may need such hydrolases to function in CS₂-rich environments. T. thioparus has been shown to consume CS₂.⁸²

3.2. Methane monooxygenase (MMO) and ammonia monooxygenase (AMO)

Carbon disulfide has been shown to be an inhibitor of nitrification in Nitrosomonas and Nitrosococcus bacterial strains^83,84 and reported to inhibit methane monooxygenase (MMO).⁸⁵ From a detoxification viewpoint, MMO is evolutionarily and mechanistically similar to ammonium monooxygenase (AMO),⁸⁵ and methanotrophs that depend on MMO and live in high sulfur environments (e.g., Methylomicrobium alcaliphilium) have active CS₂ hydrolases. The underlying mechanism proposed for CS₂ inhibition of nitrification involves reaction at a nucleophilic amine (lysine) or thiol (cysteine) to form a DTC or TTC near the active site of AMO.^84,86 The resulting 1,1-dithiolate may then chelate and sequester the active site copper or modify the activity owing to structural or electronic changes. An alternative mechanism would be reversible CS₂ insertion into metal–N or –S bonds as discussed above. Thus, one can envision CS₂ diffusing to a reactive site and either inserting in a metal–ligand bond motif or binding a proximal amine or thiolate to form a protein bound TTC or DTC as illustrated in pathways i and ii, respectively, in Fig. 2. In either case, the resulting changes in structure, electronics, metal chelation, etc. would impact protein function.

Fig. 2 Possible interactions of CS₂ with biological metals. Since it is lipophilic, CS₂ could readily transport to the active sites of cellular proteins and (i) insert into a metal–cysteine bond to form a trithiocarbonate chelate or (ii) bind a protein amine or thiol proximal to a metal center where the resulting dithiocarbamate or trithiocarbonate can bind. X = NR′ or S.

3.3. Carbon monoxide dehydrogenase (CODH)

CODH interconverts CO to CO₂ in anaerobic bacteria and is responsible for the final step of the production of acetyl coenzyme A (acetyl-CoA). Ragsdale and coworkers⁸⁷ have reported that CS₂ is a rapid binding, reversible inhibitor of this Ni/Fe–S cluster protein. Consistent with its interaction with CA, CS₂ does not bind or interact with the CO₂-binding site. Instead it serves as a competitive inhibitor to the terminal binding site of CO at an iron center of the cluster, the site of acetyl-CoA synthesis. Two proposals were offered regarding CS₂ binding at this site: insertion into a ligand–iron bond or coordination as CS₂ to the metal. An alternative possibility would be CS₂ binding via electrophilic reaction at a protein thiol or amine near the coordination site. Equally likely is that CS₂ binds the thiol of acetyl-CoA itself. Since CODH catalyzes the acetylation of the homocysteine thiol moiety of CoA,⁸⁸ CS₂ binding at this thiol would generate a TTC that inhibits CODH via chelation of the iron.

3.4. Potential targets in mammalian physiology

In general, CS₂ exposure in humans and assorted mammals has been assessed via detection of the metabolites thiazolidine-2-thione-4-carboxylic acid (TTCA), 2-thio-5-thiazolidinones, and occasionally 2-oxothiozolidine-4-carboxylic acid (Fig. 3) in the urine.^{27,30,32,89–91} TTCA is the product of CS₂ reaction with cysteine or cysteine-containing poly/oligopeptides, and its urinary release has been shown to increase linearly with CS₂ oral intake by rats.⁹² Other major metabolites, such as the 2-thio-5-thiazolidinones, likely result from terminal amines in polypeptides and other biological amines acting as nucleophiles at the CS₂ carbon to form (initially) DTCs followed by eventual cyclization. These metabolic scenarios are considered below.

Fig. 3 Cyclic metabolites found in the urine of mammals after exposure to CS₂.

3.4.1. Thiols and trithiocarbonates (TTCs). Biological thiols such as glutathione (GSH) and cysteine are essential for maintaining oxidative homeostasis in mammalian cells and serum. GSH and its oxidized partner GSSG constitute a redox buffer system that helps mitigate cellular damage from reactive radicals. GSH is present in cytosols at millimolar concentrations,⁹³ and while in plasma these lower weight thiols are present in very low concentrations (0.1 to 20 μM), plasma-based protein sulfhydryls are as high as 0.5 mM.^94,95 Owing partially to their high concentrations and CS₂'s electrophilicity, such thiols must be considered likely targets or biological sinks for CS₂. This interaction will strongly depend on the pK_a of the specific thiol and on the local pH. Biological sulfhydryls (R–SH) can display reasonably low pK_a values, though these are medium dependent. The pK_a of GSH can be as high as 9.2 (extrapolated to zero ionic strength)^96–98 and as low as around 8.66.⁹⁹ Similarly, the pK_a of free cysteine ranges from 8.2¹⁰⁰ to 8.6⁹⁷ and varies considerably within proteins. Peptide amines are somewhat more basic (pK_a ≥ 9) than these thiolates and when protonated will not be nucleophiles for CS₂. Thiols and thiolates (R–S⁻), therefore, remain the predominant nucleophiles in biological media.¹⁰¹ This should lead to the reversible formation of TTCs as illustrated in the first step of Scheme 2 and in eqn (6).

Scheme 2 Pathway for the formation of TTCA from the reaction of GSH with CS₂.^30,49

As aforementioned, TTCA release through the urinary tract is a signature of external exposure to CS₂. This thiazolidine is apparently derived from the reaction of CS₂ with the thiolate of cysteine or GSH, in the latter case resulting also in liberation of glycine and L-glutamic acid.^89,102,103 However, it is not known whether TTCA formation is enzyme assisted. While reaction with a secondary amine of GSH to give a DTC is a mechanistic possibility, such sites are much less nucleophilic than the thiolate.¹⁰⁴ In either case, intramolecular cyclization and liberation of H₂S would result in TTCA as roughly outlined in Scheme 2.

The formation of alkyl TTCs in aqueous solution is reversible (eqn (6)),¹⁰⁵ although the equilibria are dependent on the pH and on the pK_a of the parent thiol. Such equilibria are not well characterized with biological thiols, but these are especially interesting given that TTCs may be a means to sequester or transport CS₂ in serum or to provide a pathway for the interactions of CS₂ with metalloproteins. In addition, TTC formation as post-translational protein modification at cysteine residues likely leads to important structural and/or functional effects that also may be important signaling pathways, even if such reactions are reversible. Protein microenvironments can markedly affect the acidity of a thiol through interactions with charged amino acid side chains and/or hydrogen bonding.^99–101 Such interactions with protein thiols are potential targets and switches for CS₂ signaling.

Another signaling mechanism may involve the direct reaction of TTCs with transition metal centers or by reversible CS₂ insertion into metal thiolate bonds to form TTC metal complexes (Fig. 2). The latter pathway was demonstrated with a variety of metal–thiolate complexes under ambient conditions^57,68,69 including the biologically relevant zinc(II) and copper(I) centers (see above). Thus, one can envision metal thiolate centers such as zinc fingers or iron redox centers as targets for the reversible formation of CS₂ adducts. Although the above examples were in non-aqueous media, similar reactivity may be relevant in the cores of particular proteins or other lipophilic areas.

3.4.2. Amines and dithiocarbamates (DTCs). DTCs and their derivatives (e.g., disulfides of the type R₂NC(S)SSC(S)NR₂) already have a number of medical applications. CS₂ spontaneously reacts with endogenous amines to form DTCs^30,106–108 as the first step in a sequence that converts free and terminal amino acids to thiazolidine-type cyclic moieties (2-thio-5-thiazolidinones)^89,109 (Scheme 3). An important feature of DTCs is the ability to chelate metal ions such as copper or zinc, often binding to them in the enzyme pocket^110–112 or potentially even stripping them from proteins.¹¹⁰ The high binding affinity to heavy metal ions finds medicinal application in chelation therapy through this latter route.^113,114 DTCs may also sequester metals essential for certain protein functions, and, thus have been explored as possible drugs targeting HIV/AIDS via inhibition of reverse transcriptase activity.^115–118

Scheme 3 Conversion of an amino acid to a DTC, followed by subsequent cyclization to 2-thio-5-thiazolidinone.

DTCs are commonly used fungicides and pesticides, and thus their toxicity has been reviewed^119,120 most recently by Rodrigues-Lima and coworkers.¹²¹ Many of the pathologies of DTC toxicity mirror those documented for CS₂, including potent central nervous system (CNS) effects under acute exposure as well as links to Parkinson's disease via induction of severe CNS depression.^122–124 DTC disulfides also have neuropathies very similar to that of those produced by CS₂ exposure,^125–129 although a different mechanism of toxicity has been proposed.¹²⁹

The similar symptoms of DTCs and CS₂ toxicity suggest a shared mechanism. Under acidic conditions, N,N-diethyldithiocarbamate (DEDTC) releases CS₂, but it is generally assumed that DEDTC is stable at or above neutral pH.^130,131 Valentine and coworkers¹⁰³ investigated possible in vivo CS₂ release from disulfiram and DTCs such as DEDTC to establish whether such release induced symptoms attributed to toxicity characteristic of CS₂ exposure. Using radiolabeling, they found a marked increase in urinary TTCA attributed to the presence of free CS₂ regardless of the compound tested (disulfiram or a DTC). Not surprisingly, oral exposure to DEDTC gave much higher fluxes of TTCA owing to the acidic environment of the dietary tract. Incorporation of the labeled carbon at the thiocarbonyl of TTCA indicates that CS₂ is generated from the various DTCs. This result may answer a question raised by Cvek and Dvorak in a review of DTC cellular interactions,¹³² namely, that it is unclear how polar DTCs might enter a cell through lipid bilayers. The in vivo study by Valentine et al.¹⁰³ supports the view that DTCs and CS₂ are in labile equilibria. This lability under physiological conditions thereby provides a mechanism to transport this functionality through hydrophobic membranes by passive diffusion of CS₂.

The reaction between neuronal amines and CS₂ has often been hypothesized to be the source of carbon disulfide-induced neuropathological effects.^30,106,107 Catecholamine neurotransmitters, including epinephrine (adrenaline), norepinephrine (noradrenaline), and dopamine have amine-bearing side chains sufficiently nucleophilic¹³³ to react with CS₂in vivo. This reaction is a likely cause of the significant norepinephrine depletion in the adrenal glands and storage granules of rodents exposed to CS₂ vapor (64 ppb).¹⁰⁶ Norepinephrine is produced from dopamine via dopamine β-hydroxylase (DBH), a monooxygenase with copper in the active site,¹⁰⁷ and it has been suggested¹⁰⁶ that DBH is inhibited by DTC formation and chelation of that copper. Experiments in vitro showed a neuronal amine is needed to inhibit DBH in the presence of CS₂ and that this effect is reversible.¹⁰⁷ Notably, CS₂ proved to be more effective inhibitor of DBH than an added glycine-based DTC.¹⁰⁷ This suggests that lipophilic CS₂ enters catecholamine storage granules where it reacts to form DTCs and inhibits DBH (Fig. 4). Such a process could play a regulatory role by controlling dopamine/norepinephrine levels.^134,135

Fig. 4 Release of CS₂ from DTCs allows for diffusion across membranes, like the storage granules of catecholamines. Dopamine β-hydroxylase (DBH), a copper-centered enzyme, converts dopamine to norepinephrine and is deactivated by CS₂via the DTC of catecholamines.¹⁰⁷

Disulfiram (also called “antabuse”), the disulfide of DEDTC, has long been used in alcohol aversion therapy.¹³¹ It functions by inhibiting acetaldehyde dehydrogenase (ALDH), thereby greatly enhancing the severity of hangovers due to alcohol consumption. The molecular mechanism of ALDH inhibition is mediated by the metabolic products of disulfiram, the first of which is DEDTC formed by reduction of the disulfide bond (Fig. 5). Metabolites of DEDTC are irreversible inhibitors of ALDH.¹²¹ Covalent addition of one of these metabolites to an essential cysteine residue inhibits rodent ALDH in vivo by initiating disulfide bond formation. The current thinking is that oxidation by microsomes forms methyl diethylthiocarbamoyl sulfoxides (Fig. 5) as the active agent.^130,136 Direct reactions with DTCs or their disulfides have also been proposed.^119,135 Either of these mechanisms provide a route for CS₂ to interact with specific proteins having ideally placed cysteines and could be essential in bio-signaling chemistry (Fig. 6).^113,136,138

Fig. 5 Disulfiram is readily reduced in vivo to form DEDTC, which generates carbon disulfide or is eventually converted to a thiocarbamoyl. All have been implicated in inhibiting ALDH via mixed disulfide formation with protein thiols.

Fig. 6 Possible interactions of CS₂-adducts with proteins suggested by the inhibitory action of disulfiram and its metabolites with ALDH. Formation of a mixed disulfide on a critical protein thiol leads to oxidation and intra-protein disulfide formation with a nearby thiol. X = NR′ or S.

Disulfiram is also finding therapeutic use in treating cocaine addiction via an ALDH-independent mechanism.^137,139 This may function by inhibiting DBH, the copper-dependent enzyme required for metabolism of dopamine to norepinephrine. These inhibitory effects have been attributed to the copper-chelating activity of DEDTC formed in vivo. Such copper-chelation is also thought to induce proteasome inhibition leading to the proposal that disulfiram could serve in anticancer therapy.¹⁴⁰

Although there is strong evidence implicating CS₂ generation in DTC toxicity,¹³⁰ it is important to consider concentration regimes. One proposed mechanism for CS₂ neuropathy involves the induction of intra- and inter-protein cross-linking. The proteins β-lactoglobulin, porcine neurofilaments, bovine serum albumin, and hemoglobin were each observed to react with CS₂ to form thiourea based crosslinks and dimers. This involved predominately lysine ε-amines,¹²⁹ but also lysine and the N-termini of certain proteins,^141,142 as the pK_a of such amines can be significantly decreased in protein microenviornments.¹⁴³ DTC formation on an exposed amine, followed by loss of HS⁻ would give isothiocyanate intermediates that couple with another free amine (Scheme 4).¹⁴⁴ However, it should be noted that these in vivo processes were demonstrated in rats exposed to 50 ppm CS₂ (5 times the occupational threshold)¹⁹ in inhaled air for 2 weeks.

Scheme 4 Transformation of CS₂ derived lysine dithiocarbamates into dimers responsible for CS₂-induced protein cross-linking via thioureas moieties.¹⁴¹

3.4.3. Nuclear factor κB (NF-κB). DTCs are known inhibitors of NF-κB,^132,145 a class of closely related, pro-inflammatory factors.^146–154 NF-κB is a crucial mediator in inflammation-based tumor growth and progression¹⁴⁶ and triggers the transcription of genes that regulate cellular proliferation and differentiation,¹⁵⁰ angiogenesis, and proinflammatory cytokines and chemokines. NF-κB directly and indirectly affects a wide range of functions from the vascular endothelium to immune response,¹⁵² thus making it an attractive therapeutic target.

What stimulates the constitutive expression of NF-κB in cancer cells is not fully understood.¹⁵⁵ An extracellular signal triggers a cytosolic-based cascade to release NF-κB from an inhibitor (I-κB). Following translocation to the nucleus, NF-κB tunes the regulation of diverse genes. To activate NF-κB, the I-κB must be phosphorylated by an I-κB kinase (IKK) resulting in its conjugation to ubiquitin (ubiquitination) and subsequent degradation.^{132,156–159} DTCs can act as thiol antioxidants, which are also known to inhibit NF-κB.¹⁵² However, the inhibitory activity of DTCs appears to be isolated from its antioxidant abilities¹⁶⁰ and relies on a mechanism that has not been fully elucidated.

A potential target for CS₂ is a cysteine (Cys38) in the DNA binding site of NF-κB (Fig. 7, pathway 1).¹⁶¹ Treatment with thiol-reactive compounds such as N-tosyl-L-phenylalanine chloromethyl ketone (TPCK) prevents DNA binding.¹⁵² TPCK reacts far more freely at this residue than it does with the cellular GSH, suggesting that the Cys38 thiol is more nucleophilic or has a lower pK_a,¹⁶¹ thereby making it a likely target for CS₂via the reversible formation of a TTC.¹⁵⁰ Such adduct formation could also provide a mechanism for DTC inhibition of NF-κB, given that dithiocarbamates release CS₂in vivo (see above).

Fig. 7 Simplified depiction of NF-κB activation and gene up-regulation to produce pro-inflammatory proteins. The IKK complex (top right) is activated through a signal cascade and in turn activates NF-κB. Possible targets for CS₂ inhibition are the Cys38 in the DNA binding region of NF-κB (pathway 1), the Cys179 in IKKβ (pathway 2), or lysines in the zinc finger binding domains of the pre-ubiquitinated scaffold protein NEMO (pathway 3). Cys179 appears to be the most ubiquitous and likely target. (IL-6 = interleukin-6, a proinflammatory cytokine, iNOS = inducible nitric oxide synthase, I-κB = inhibitor of NF-κB, NEMO = NF-κB essential modulator (also known as IKKγ), IKK = I-κB kinase).^{146,148,152,155,159,161–164}

Thiolate-reactive compounds also inhibit the IKK complex further upstream in the signaling cascade (Fig. 7, pathway 2). The presumed target is again a critical cysteine (Cys179), located between two serine residues (Ser177/181) in the activation loop of the protein β-subunit.¹⁶⁴ Ser177/181 are each phosphorylated by another kinase to activate IKK (Fig. 7).¹⁶⁵ Replacing Cys179 with alanine prevents phosphorylation, indicating that this cysteine is non-innocent in the activation step.¹⁶⁶ Further, Cys179 is implicated as a participant in the catalytic domain of activated IKK^164,167 due to significant (albeit reversible) enzyme deactivation occurring with DTC treatment of pre-phosphorylated IKKβ. The fact that DTC inhibition is reversible suggests a mechanism distinct from the disulfide formation step that is responsible for ALDH inhibition. Indeed, CS₂ liberated from a dithiocarbamate may form a TTC at Cys179, thereby providing a reversible IKK inhibition pathway.

Another possible upstream target for CS₂ is the NF-κB essential modulator (NEMO), which recruits other IKK activators (Fig. 7, pathway 3). NEMO can be activated by ubiquitination at lysine residues including those in a zinc finger binding domain.^159,162 This suggests another site for CS₂/DTC interference. DTCs have been shown to decrease the expression of NEMO¹⁵⁵ in addition to the IKKβ, consistent with this hypothesis.

NF-κB is an alluring therapeutic target due to the prevalence of heightened inflammation in a variety of disease states.^70,132,152 If CS₂ is indeed an SMB that plays a critical role in the regulation of these important transcription factors, then it stands to reason that CS₂ delivery may have therapeutic value, notably in the context of inflammation reduction. Along this line of thought, delivery of CS₂ has been proposed in the patent literature for the treatment of inflammatory conditions and was claimed to directly inhibit NF-κB.¹⁶⁸

3.4.4. Interactions with cytochrome P450. The hindered ability to metabolize drugs oxidatively is a symptom also attributed to CS₂ toxicity at relatively high levels of exposure,³⁰ and this suggests inhibition of hepatic monooxygenases such as cytochrome P450. The interaction of ¹⁴CS₂ with P450, NADPH, and O₂ results in the expiration of some ¹⁴CO₂,¹⁶⁹ and since carbonic anhydrases are inactive for CS₂ hydrolysis (Section 3.1), another mechanism needs to be considered. Oxidative desulfuration^169,170 is proposed to occur via CS₂ oxidation to an unstable S-oxygenated intermediate¹⁷¹ that fragments to elemental sulfur and to COS or monothiocarbonate.^90,172 Carbonic anhydrase can hydrolyze COS to H₂S and CO₂, while monothiocarbonate reacts with carbamyl phosphate synthetase to form thiourea.³¹ The zero-valent byproduct S(0) can modify activities of the targeted proteins by forming persulfides with cysteine residues.^171,173,174 This modification may be one cause of hepatic and renal toxicity,¹⁰⁸ although such toxicity has also been attributed to the H₂S from CA catalyzed COS hydrolysis.¹⁷² Given the bioregulatory properties of H₂S, such release may have other physiological implications.^163,175 Other potential CS₂-derived sources of H₂S include TTC cyclization and DTC conversion to an isothiocyanate.

3.4.5. Persulfides and selenoproteins. To this point we have largely focused on the reactions of CS₂ with the more common biological amine and thiol nucleophiles. Reactions with thiols (or more appropriately thiolates) are readily predicted, since these are among the most prevalent and potent nucleophiles present and are capable of reacting with both “hard” and “soft” electrophiles.¹⁷⁶ However, the recent discovery of ubiquitous and significant levels of hydropersulfides (RSSH) in mammalian cells, tissue and plasma^177,178 presents other likely targets for CS₂ reactivity/biological activity. Numerous protein persulfides have been detected and the concentration of glutathione hydropersulfide (GSSH) has been found to be as high as 150 μM in mouse brain and 50 μM in mouse liver and heart, although their biological functions are still being elucidated. These levels make GSSH the second most prevalent sulfur species in cells, behind only GSH itself. The possible reaction between CS₂ and hydropersulfides seems especially relevant, since hydropersulfides are much more reducing and nucleophilic than the analogous thiols.^179,180 These properties predict a more efficient and favorable reaction with persulfides compared to thiols. Moreover, hydropersulfides are more acidic than the corresponding thiols by about 1–2 pK_a units, indicating a higher percentage of the nucleophilic persulfide anion compared to thiolates at physiological pH. To our knowledge, the reaction between hydropersulfides and CS₂ has not been reported, but it is not difficult to envision formation of perthioxanthates (RSSCS₂⁻) (eqn (11)).

Another potent class of biological nucleophiles is represented by selenoproteins. Like hydropersulfides, selenocysteine is more acidic than cysteine by 2–3 pK_a units. Thus, selenocysteines exist predominantly as the anionic selenide at physiological pH. In general, selenols are stronger nucleophiles than are thiols under conditions of equal ionization and even more so under physiological conditions where a higher percentage of the selenide anion would be present compared to thiolate.¹⁸¹ Of course, selenoproteins are far less prevalent compared to thiol proteins. To date, only about 25 human selenoproteins have been identified with functions ranging from thyroid hormone regulation to antioxidant function.¹⁸² For comparison, the human genome encodes for over 200 000 cysteines that are widely distributed among proteins.¹⁸³ The reaction of a selenide with CS₂ can be envisioned to be analogous to that shown for the thiol/thiolate with the generation of the selenoxanthate (RSeC(S)S⁻). Therefore, due to the nucleophilicities of persulfides and selenols, future studies into the chemical biology of CS₂ should consider the possible interactions with persulfides and/or selenoproteins.

3.5. Endogenous production?

A hallmark of the SMBs NO, CO, and H₂S is that each is generated endogenously by enzymatic processes.¹ While at present there is no direct evidence for mammalian CS₂ production by a specific mammalian enzyme, there is ancillary evidence for endogenous production.^184,185 For example, humans (controlled for exposure, medication, etc.) exhale CS₂. While healthy individuals do not exhale it at levels where there is a significant difference in alveolar gradient,¹⁸⁶ elevated CS₂ levels on the breath are associated with a variety of disease states.¹⁸⁵ For instance, patients with schizophrenia have high positive CS₂ gradients, indicating higher CS₂ levels in their blood steams.¹⁸⁷ Given CS₂'s interaction with catecholamines resulting in increased dopamine levels,¹⁰⁷ and the association of high levels of dopamine with schizophrenia,^134,135 an abundance of CS₂ may play a role. Exhaled CS₂ is also associated with gastric cancers, especially those induced by Helicobacter pylori.¹⁸⁴ A significant increase in CS₂ is found in human feces after eating, suggesting a role of the gut microbiome in its production.^188,189 Moreover, it is found in all samples of healthy donors and missing in many patients with foodborne bacterial infections like Campylobacter jejuni,¹⁸⁸ implicating a positive association of CS₂ production with a healthy gut microbiome.

Exhaled CS₂ is also claimed to be a marker for organ rejection after lung transplantation,^184,185 although COS has also been claimed as such a marker.¹⁹⁰ It is notable that CS₂ or COS would be associated with organ transplantation, as it is well-established that CO has protective effects in organ transplantation, reducing probability of rejection.⁴

What are the sources of exhaled CS₂? It is possible that, like other sulfur containing volatiles, it is a by-product of metabolic pathways of methionine.^184,185 Alternatively, it has been inferred that it derives from a collection of gut bacteria and micro-organisms of the human microbiome that produces other sulfur-containing small metabolites.^187–189 However, one should also consider the possibility of a dedicated, enzymatic source. This is most apparent in certain rodents that exhale CS₂ in micromolar concentrations.¹⁹¹ The olfactory system of such rodents appears to have sensory neurons sensitized to CS₂ through an isoform of membrane-bound receptor guanylyl cyclase, type D (GC-D).¹⁹² This receptor triggers a cyclic guanosine monophosphate (cGMP)-signaling cascade, much like the response of sGC to NO. This CS₂-initiated cascade, when combined with an odor, promotes in rodents a preference for the food associated with that odor, an important instinctual survival mechanism.¹⁹¹ Knockouts for the gene encoding the olfactory GC-D resulted in loss of the ability to form these preferences.^192,193 This result implies that, at least with these rodents, CS₂ has a signaling-inducing target rather than being simply a metabolic byproduct. A form of CA II has also been reported to play a role in such signal transduction,¹⁹² even though other CAs generally do not interact strongly with CS₂.⁷⁵ The murine olfactory system is also sensitive to CO₂, but by several orders of magnitude less than to CS₂. These observations suggest that CS₂ is endogenously produced in mammals at concentration regimes appropriate for an SMB.

3.5.1. Bodily inventory of CS₂. A quantitative relationship between the CS₂ present in humans (generally assumed to be from external exposure) and temporal parameters is challenging to establish.^30,194,195 Few studies account for chronic exposure¹⁹⁶ or the limits of quantification. CS₂ exposure has largely been assessed via detection of the urinary metabolite TTCA (Scheme 4). Occupational atmospheric exposure limits are set as low as 5 ppm, a level that generates detectable quantities of TTCA³⁸ but does not show conspicuous negative health effects. Various sources have found that TTCA is not detected without external exposure, and therefore have argued against endogenous production.^197–199 The TTCA detection limit is ∼1 μM in urine, which correlates approximately with atmospheric exposure of 0.2–0.4 ppm CS₂.^195,197 Other studies have estimated that only 2–6% of CS₂ ingested is metabolized to TTCA^89,197–202 owing to alternative metabolic pathways as well as loss through respiration. Thus, one can argue that low endogenous levels of CS₂ may go undetected. For comparison, NO has dynamic and contrasting functions over concentrations from as low as a few nanomolar for vascular functions, to 100 nM for angiogenic and anti-apoptotic effects, to 300 nM for apoptotic conditions, to a few micromolar under nitrosative stress.²⁰³

Notably, TTCA is detected in the urine of individuals not directly exposed to CS₂.²⁰⁴ Specifically, consumption of brassica vegetables such as broccoli, cauliflower, and cabbage results in urine excretions of 10 μmol TTCA per liter of urine. This observation suggests that these vegetables may be a natural source of CS₂ given the nature of TTCA formation.¹⁷ While TTCA is also found in the vegetables themselves, far greater quantities are found when they are boiled.²⁰⁵ Given that TTCA is derived from the reaction of GSH or cysteine with CS₂, it is reasonable that heat simply increases the rate of this reaction and thus of TTCA production. Hence, it stands to reason that these food sources may contain trace amounts of CS₂ or a precursor thereof. Moreover, brassica vegetables are promoted as healthy and beneficial foods due to their association with lower cancer risk in a multitude of studies.²⁰⁶ The bioactive compounds are the sulfur containing phytochemicals glucosinolates (Fig. 8). They have anti-inflammatory, genetic, and epigenetic properties once metabolized and have been probed for therapeutic potential. Glucosinolates undergo enzymatic conversion to form isothiocyanates and DTCs, which in turn can generate CS₂ and TTCA.^205,207

Fig. 8 Brassica vegetables contain glucosinolates, which undergo enzymatic conversion to a variety of metabolites, with various effects, including anti-inflammatory action. Since DTCs are generated, these compounds may also yield CS₂.

4. Summary and outlook

In this review, we've attempted to draw together literature evidence indicating that carbon disulfide may have roles in mammalian biology beyond its toxicity. The various chemical pathways of CS₂ that might contribute to such properties are summarized in Fig. 9. We have posed the questions: does CS₂, its precursors, or the resulting metabolites have therapeutic potential? and/or Is CS₂ an unrecognized small molecule bioregulator? Like NO, CO, and H₂S, carbon disulfide is a small molecule that is nonpolar and diffusible and is also known to be toxic at relatively high concentrations. Furthermore, there is fragmentary evidence that CS₂ is produced endogenously, even in humans, although it is possible that this production is at least in part from the closely associated microbiome. The biological chemistry and metabolism of CS₂ suggest a commonality in biochemical targets with other SMBs. This includes reactivity with amino acids like cysteine and lysine and the formation of adducts in cytosol, serum, and proteins. Metal centers are also likely targets via the insertion of CS₂ into metal ligand bonds to form chelating ligands. These reactions readily occur at ambient temperatures and in aqueous media. Such pathways may initiate interconnected signaling cascades. For examples, reactions of CS₂ with biological nucleophiles in some cases release H₂S, and the inhibition of NF-κB is critically involved in inducible nitric oxide synthase activation. Furthermore, it is entirely likely that CS₂ generated by a specific enzyme, system, etc. would be ultimately consumed and metabolized in a specific manner when functioning normally, e.g., through oxidative desulfurization in the liver.

Fig. 9 Hypothetical metabolic map for CS₂. Species highlighted in grey are known human urinary metabolites of CS₂.^{30,90,171–175} The signaling chemistry is proposed to be through CS₂ reactions with biological nucleophiles. Dotted arrows represent possible, albeit minor, pathways. H₂S (HS⁻ under physiological conditions) is produced at several sites, suggesting an interplay between CS₂ and this SMB. (SAM = S-adenosyl methionine methyltransferases, CA = carbonic anhydrases, P450 = cytochrome P450 monooxygenase, DBH = dopamine β-hydroxylase, TTCA = thiazolidine-2-thione-4-carboxylate.)

Major advances in the investigation of the bioregulatory, therapeutic, etc. roles of SMBs involved the development of strategies for controlled delivery. For example, researchers into the chemical biology of NO have long drawn on the availability of water-soluble salts of various “NONOate” anions, R₂NN(O)NO⁻, that allow for in vitro and in vivo NO generation with well-defined half-lives ranging from minutes to hours.²⁰⁸ Ultimately, analogous donors with predictable kinetics for CS₂ release under physiological conditions will need to be utilized to evince CS₂ interactions with specific biological targets. In this context, one can envision designing DTC or TTC compounds (or their metal complexes) that, given their reversible equilibria with amines or thiols, would release controlled quantities of CS₂ over time to in vitro or in vivo targets. An alternative strategy would be to use photochemical methods to uncage CS₂ as is being explored extensively with NO and CO.²⁰⁹ The advantage of the latter strategy would be the ability to control location, timing, and dosage of such release.

The observations discussed herein by no means establish an unambiguous case for the bioregulatory or therapeutic roles of carbon disulfide. But they do point to the exciting potential represented by the chemical biology of CS₂ and the relatively unexplored research challenges and opportunities presented by these possibilities.

Author contributions

AWD, DFZ, and PCF each contributed to the preparation and editing of the full manuscript. AWD prepared the initial draft and all the figures and PCF completed the final draft. JMF critically read the manuscript and contributed Section 3.4.5.

Financial interest

None of the authors have competing financial interests relevant to the contents of this article.

Abbreviations

AIDS	Acquired immunodeficiency syndrome
ALDH	Acetaldehyde dehydrogenase
AMO	Ammonium monooxygenase
CA	Carbonic anhydrase
cGMP	Cyclic guanosine monophosphate
CNS	Central nervous system
CO	Carbon monoxide
CO2	Carbon dioxide
CoA	Coenzyme A
CODH	Carbon monoxide dehydrogenase
COS	Carbonyl sulfide
CS2	Carbon disulfide
Cys	Cysteine
DBH	Dopamine β-hydroxylase
DEDTC	N,N-Diethyldithiocarbamate
DTC	Dithiocarbamate
GC-D	Guanylyl cyclase, isoform D
Glu	L-Glutamic acid
Gly	Glycine
GSH	Reduced glutathione
GSSG	Oxidized glutathione dimer
H2S	Hydrogen sulfide
HIV	Human immunodeficiency virus
I-κB	Inhibitor NF-κB
IKK (α,β)	I-κB kinase (α,β-subunits)
IL-6	Interleukin-6
iNOS	Inducible nitric oxide synthase
Lys	L-lysine
MMO	Methane monooxygenase
NAD(P)H	Reduced nicotinamide adenine dinucleotide (phosphate)
NEMO	NF-κB essential modulator (also IKKγ)
NF-κB	Nuclear factor κB
NO	Nitric oxide
P450	Cytochrome P450
SAM	S-Adenosyl methionine transmethylase
Ser	L-Serine
sGC	Soluble guanylyl cyclase
SMB	Small molecule bioregulator
TPCK	N-Tosyl-L-phenylalanine chloromethyl ketone
TTC	Trithiocarbonate
TTCA	Thiazolidine-2-thione-4-carboxylic acid

Acknowledgements

We thank Drs Maykon Lima Souza and Mary E. Howe-Grant for their careful reading of the manuscript and suggestions. Studies in the Ford laboratory on SMBs have long been supported by the Chemistry Division of the National Science Foundation.

References

1. J. M. Fukuto, S. J. Carrington, D. J. Tantillo, J. G. Harrison, L. J. Ignarro, B. A. Freeman, A. Chen and D. A. Wink, Small molecule signaling agents: the integrated chemistry and biochemistry of nitrogen oxides, oxides of carbon, dioxygen, hydrogen sulfide, and their derived species, Chem. Res. Toxicol., 2012, 25, 769–793.
2. E. Buys and P. Sips, New insights into the role of soluble guanylate cyclase in blood pressure regulation, Curr. Opin. Nephrol. Hypertens., 2014, 23, 135–142.
3. C. Szabo, Gasotransmitters in cancer: from pathophysiology to experimental therapy, Nat. Rev. Drug Discovery, 2016, 15, 185–203.
4. S. W. Ryter and A. M. K. Choi, Targeting heme oxygenase-1 and carbon monoxide for therapeutic modulation of inflammation, Transl. Res., 2016, 167, 7–34.
5. K. R. Olson, The therapeutic potential of hydrogen sulfide: separating hype from hope, Am. J. Physiol.: Regul., Integr. Comp. Physiol., 2011, 301, R297–R312.
6. I. Andreadou, E. K. Iliodromitis, T. Rassaf, R. Schulz, A. Papapetropoulos and P. Ferdinandy, The role of gasotransmitters NO, H₂S and CO in myocardial ischaemia/reperfusion injury and cardioprotection by preconditioning, postconditioning and remote conditioning, Br. J. Pharmacol., 2015, 172, 1587–1606.
7. Nitric Oxide: Biology and Pathobiology, ed. L. J. Ignarro, Elsevier Inc., Burlington MA, 2nd edn, 2010.
8. S. H. Heinemann, T. Hoshi, M. Westerhausen and S. Schiller, Carbon monoxide – physiology, detection and controlled release, Chem. Commun., 2014, 50, 3644–3660.
9. D. A. Wink, L. A. Ridnour, S. P. Hussain and C. C. Harris, The reemergence of nitric oxide and cancer, Nitric Oxide, 2008, 19, 65–67.
10. D. Wu, W. Si, M. Wang, S. Lv, A. Ji and Y. Li, Hydrogen sulfide in cancer: friend or foe?, Nitric Oxide, 2015, 50, 38–45.
11. J. Muntané and B. Bonavida, Special collection: nitric oxide in cancer, Redox Biol., 2015, 6, 505–506.
12. J. R. Lancaster Jr., A Tutorial on the diffusibility and reactivity of free nitric oxide, Nitric Oxide, 1997, 1, 18–30.
13. C. M. Bernt, P. T. Burks, A. W. DeMartino, A. E. Pierri, E. S. Levy, D. F. Zigler and P. C. Ford, Photocatalytic carbon disulfide production via charge transfer quenching of quantum dots, J. Am. Chem. Soc., 2014, 136, 2192–2195.
14. R. W. McKee, Solubility of carbon disulfide vapor in body fluids and tissues, J. Ind. Hyg. Toxicol., 1941, 23, 484–489.
15. G. L. Gutsev and R. J. Bartlett, Electron affinities of CO₂, OCS, and CS₂, J. Chem. Phys., 1998, 108, 6756–6762.
16. R. Sander, Compilation of Henry's Law constants (version 4) for water as solvent, Atmos. Chem. Phys., 2015, 15, 4399–4981.
17. S. F. Watts, The mass budgets of carbonyl sulfide, dimethyl sulfide, carbon disulfide and hydrogen sulfide, Atmos. Environ., 2000, 34, 761–779.
18. N. Rojo, G. Gallastegi, A. Barona, L. Gurtubay, G. Ibarra-Berastegi and A. Elías, Biotechnology as an alternative for carbon disulfide treatment in air pollution control, Environ. Rev., 2010, 18, 321–332.
19. WHO Concise International Chemical Assessment Document (CICAD) 46: Carbon Disulfide, World Health Organization, United Nations Environment Program, International Labor Organization, 2002.
20. H. P. Gelbke, T. Göen, M. Mäurer and S. I. Sulsky, A review of health effects of carbon disulfide in viscose industry and a proposal for an occupational exposure limit, Crit. Rev. Toxicol., 2009, 39, 1–126.
21. A. Delpech, Note sur les accidents que dévelope, chez les ouvriers en caoutchouc, l'inhalation du sulfure de carbone en vapeur, Académie Impériale de Médecine, 1856, vol. 21, p. 350.
22. S. G. Selevan, R. Hornung, J. Fajen and C. Cottrill, Paternal exposure to carbon disulfide and spouse's pregnancy experience, Department of Health and Human Services, Centers for Disease Control, NIOSH, Hazard Evaluations and Field Studies, Cincinnati OH, 1983.
23. A. M. Cirla, P. A. Bertazzi, M. Tomasini, A. Villa, C. Graziano, R. Invernizzi and R. Gilioli, Study of endocrinological functions and sexual behavior in carbon disulfide workers, Med. Lav., 1978, 69, 118–129.
24. G. Wagar, M. Tolonen and E. Helpio, Endocrinological studies of Finnish viscose rayon workers exposed to carbon disulfide, G. Ital. Med. Lav., 1981, 3, 103–105.
25. S. Y. Zhou, Y. X. Liang, Z. Q. Chen and Y. L. Wang, Effects of occupational exposure to low-level carbon disulfide (CS₂) on menstruation and pregnancy, Ind. Health, 1988, 26, 203–214.
26. J. Y. Ma, J. J. Ji, Q. Ding, W. D. Liu, S. Q. Wang, N. Wang and G. Y. Chen, The effects of carbon disulfide on male sexual function and semen quality, Toxicol. Ind. Health, 2010, 26, 375–382.
27. D. G. Graham, V. Amarnath, W. M. Valentine, S. J. Pyle and D. C. Anthony, Pathogenetic studies of hexane and carbon disulfide neurotoxicity, Crit. Rev. Toxicol., 1995, 25, 91–112.
28. A. E. Cohen, L. D. Scheel, J. F. Kopp, F. R. Stockell, R. G. Keenan, J. T. Mountain and H. J. Paulus, Biochemical mechanisms in chronic carbon disulfide poisoning, Am. Ind. Hyg. Assoc. J., 1959, 20, 303–323.
29. B. Soucek, Transformations of carbon disulfide in organisms, J. Hyg., Epidemiol., Microbiol., Immunol., 1957, 1, 10–22.
30. R. O. Beauchamp, J. S. Bus, J. A. Popp, C. J. Boreiko, L. Goldberg and M. J. McKenna, A critical review of the literature on carbon disulfide toxicity, Crit. Rev. Toxicol., 1983, 11, 169–278.
31. M. Davidson and M. Feinleib, Carbon disulfide poisoning: A review, Am. Heart J., 1972, 83, 100–114.
32. H. Drexler, T. Göen, J. Angerer, S. Abou-el-ela and G. Lehnert, Carbon disulphide, Int. Arch. Occup. Environ. Health, 1994, 65, 359–365.
33. F. Song, S. Yu, X. Zhau, C. Zhang and K. Xie, Carbon disulfide-induced changes in cytoskeleton protein content of rat cerebral cortex, Neurochem. Res., 2006, 31, 71–79.
34. S. Krstev, B. Perunicic, B. Farkic and R. Banicevic, Neuropsychiatric effects in workers with occupational exposure to carbon disulfide, J. Occup. Health, 2003, 45, 81–87.
35. T. Takebayashi, Y. Nishiwaki, T. Nomiyama, T. Uemura, T. Yamauchi, S. Tanaka, H. Sakurai and K. Omae, Lack of relationship between occupational exposure to carbon disulfide and endocrine dysfunction: a six-year cohort study of the Japanese rayon workers, J. Occup. Health, 2003, 45, 111–118.
36. C. Meyer, Semen quality in workers exposed to carbon disulfide compared to a control group from the same plant, J. Occup. Med., 1981, 23, 435–439.
37. A. M. Cirla and C. Graziano, Health impairment in viscose-rayon workers with carbon disulfide risk below 30 mg m⁻³. An exposed-controls study, G. Ital. Med. Lav., 1981, 3, 69–73.
38. J. Domergue, D. Lison and V. Haufroid, No evidence of cardiovascular toxicity in workers exposed below 5 ppm carbon disulfide, Int. Arch. Occup. Environ. Health, 2016, 89, 835–845.
39. A. Schramm, W. Uter, M. Brandt, T. Goen, M. Kohrmann, T. Baumeister and H. Drexler, Increased intima-media thickness in rayon workers after long-term exposure to carbon disulfide, Int. Arch. Occup. Environ. Health, 2016, 89, 513–519.
40. T. O. Peyton, R. V. Steel and W. R. Mabey, Carbon disulfide, carbonyl sulfide: literature review and environmental assessment, Stanford Res. Inst., 1976.
41. S. R. Rao, Xanthates and Related Compounds, Marcel Dekker, Inc., New York City NY, 1971.
42. W. Zeise, Vorleufige Anzeige einer neuen Klasse von Schwefelverbindungen, J. Chem. Phys., 1822, 35, 175–182.
43. S. R. Rao and C. C. Patel, Kinetics of decomposition of xanthates, J. Mines, Met. Fuels, 1960, 8, 9–14.
44. E. Humeres, N. A. Debacher, M. M. de S. Sierra, J. D. Franco and A. Schutz, Mechanisms of acid of dithiocarbamates. 1. Alkyl dithiocarbamates, J. Org. Chem., 1998, 63, 1598–1603.
45. E. Humeres, N. A. Debacher and M. M. de S. Sierra, Mechanisms of acid decomposition of dithiocarbamates. 2. Efficiency of the intramolecular general acid catalysis, J. Org. Chem., 1999, 64, 1807–1813.
46. E. Humeres, B. Sun Lee and N. A. Debacher, Mechanisms of acid decomposition of dithiocarbamates. 5. Piperidyl dithiocarbamate and analogues, J. Org. Chem., 2008, 73, 7189–7196.
47. D. De Filippo, P. Deplano, F. Devillanova, E. F. Trogu and G. Verani, Inductive effect in dithiocarbamate decomposition mechanism, J. Org. Chem., 1973, 38, 560–563.
48. W. O. Foye, J. R. Marshall and J. Mickles, Structure of the carbon disulfide adduct of β-mercaptoethylamine, J. Pharm. Sci., 1963, 52, 406–407.
49. T. Weiss, J. Hardt and J. Angerer, Determination of urinary 2-thiazolidinethione-4-carboxylic acid after exposure to alkylene bisdithiocarbamates using gas chromatography–mass spectrometry, J. Chromatogr. B: Biomed. Sci. Appl., 1999, 726, 85–94.
50. B. Holmberg, Über Thiocarbonate, J. Prakt. Chem., 1906, 73, 239–248.
51. K. K. Pandey, Reactivities of carbonyl sulfide (COS), carbon disulfide (CS₂) and carbon dioxide (CO₂) with transition metal complexes, Coord. Chem. Rev., 1995, 140, 37–114.
52. J. A. Ibers, Centenary Lecture. Reactivities of carbon disulphide, carbon dioxide, and carbonyl sulphide towards some transition-metal systems, Chem. Soc. Rev., 1982, 11, 57–73.
53. M. M. Doeff, Reaction of matrix-isolated iron atoms with carbon disulfide, Inorg. Chem., 1986, 25, 2474–2476.
54. H. Werner, Novel Coordination Compounds Formed from CS₂ and heteroallenes, Coord. Chem. Rev., 1982, 43, 165–185.
55. A. Bheemaraju, J. W. Beattie, R. L. Lord, P. D. Martin and S. Groysman, Carbon disulfide binding at dinuclear and mononuclear nickel complexes ligated by a redox-active ligand: iminopyridine serving as an accumulator of redox equivalents for the activation of heteroallenes, Chem. Commun., 2012, 48, 9595–9597.
56. T. K. Muraoka, A. P. Duarte, P. M. Takahashi, V. M. Nogueira, A. V. G. Netto and A. E. Mauro, Reactivity of Fe(CO)₂(η²-CS₂)(PPh₃)₂ towards copper(I) compounds: synthesis of binuclear heterometallic species having CS₂ bridging Fe(0) and Cu(I), Eclética Quím., 2007, 32, 7–11.
57. K. Tang, X. Jin and Y. Tang, Insertion of CS₂ into M–S bonds and its Application in Synthesis of Clusters, Rev. Heteroat. Chem., 1996, 15, 83–114.
58. I. K. Bezougli, A. Bashall, M. McPartlin and D. M. P. Mingos, Insertion of CS₂ into the Group 2 metal-alkoxide bonds of [{M(OR)₂}_n] (M = Mg, Ca, Sr, or Ba; R = Et or Prⁱ), J. Chem. Soc., Dalton Trans., 1998, 2671–2678.
59. K. H. Yih, S. C. Chen, Y. C. Lin, G. H. Lee and Y. Wang, New syntheses of [Et₄N][(C₅H₁₀NCS₂)M(CO)₄] (M = Cr, Mo and W); Insertion reaction of carbon disulfide into the metal-nitrogen bond, J. Organomet. Chem., 1995, 494, 149–155.
60. N. J. Hartmann, G. Wu and T. W. Hayton, Activation of CS₂ by a masked terminal nickel sulfide, J. Chem. Soc., Dalton Trans., 2016 10.1039/C6DT00885B.
61. K. B. Ghiassi, D. T. Walters, M. M. Aristov, N. D. Loewen, L. A. Berben, M. Rivera, M. M. Olmstead and A. L. Balch, Formation of a Stable Complex, RuCl₂(S₂CPPh₃)(PPh₃)₂, containing an unstable zwitterion from the reaction of RuCl₂(PPh₃)₃ with carbon disulfide, Inorg. Chem., 2015, 54, 4565–4573.
62. A. C. Lane, M. V. Vollmer, C. H. Laber, D. Y. Melgarejo, G. M. Chiarella, J. P. Fackler, X. Yang, G. A. Baker and J. R. Walensky, Multinuclear copper(I) and silver(I) amidinate complexes: synthesis, luminescence, and CS₂ insertion reactivity, Inorg. Chem., 2014, 53, 11357–11366.
63. B. J. McCormick and R. I. Kaplan, Reaction of carbon disulfide with nickel(II)–amine complexes, Can. J. Chem., 1970, 48, 1876–1880.
64. J. M. Andrews, D. Coucouvanis and J. P. Fackler, Thioxanthate complexes. Carbon disulfide eliminations to form bridged mercaptide dimers of nickel(II) and palladium(II), Inorg. Chem., 1972, 11, 493–499.
65. A. H. Ewald and E. Sinn, Magnetically anomalous thio complexes of iron(III) and nickel(II), Aust. J. Chem., 1968, 21, 927–938.
66. D. Coucouvanis, S. J. Lippard and J. A. Zubieta, Sulfur-bridged dimeric complexes of iron(III), J. Am. Chem. Soc., 1970, 92, 3342–3347.
67. D. Coucouvanis, S. J. Lippard and J. A. Zubieta, Dimeric iron(III) complex containing two ethyl mercaptide and two ethyl thioxanthate groups as bridging ligands, J. Am. Chem. Soc., 1969, 91, 761–762.
68. H. E. Mabrouk and D. G. Tuck, The direct electrochemical synthesis of zinc and cadmium derivatives of α,ω-alkanedithiols and their reaction with carbon disulphide, Inorg. Chim. Acta, 1988, 145, 237–241.
69. A. Avdeef and J. P. Fackler Jr., Bis(triphenylphosphine)ethyltrithiocarbonatocopper(I), J. Coord. Chem., 1975, 4, 211–215.
70. S. M. Schmitt, M. Frezza and Q. P. Dou, New applications of old metal-binding drugs in the treatment of human cancer, Front. Biosci., Scholar Ed., 2012, S4, 375–391.
71. E. R. T. Tiekink and I. Haiduc, Stereochemical aspects of metal xanthate complexes: molecular structures and supramolecular self-assembly, Progress in Inorganic Chemistry, John Wiley & Sons, Inc., 2005, pp. 127–319.
72. E. R. T. Tiekink and J. Zukerman-Schpector, Stereochemical activity of lone pairs of electrons and supramolecular aggregation patterns based on secondary interactions involving tellurium in its 1,1-dithiolate structures, Coord. Chem. Rev., 2010, 254, 46–76.
73. J. N. Verpeaux, M. H. Desbois, A. Madonik, C. Amatore and D. Astruc, Electron-transfer-catalyzed chelation of dithiocarbamate iron complexes [Fe(η⁵-C₅R₅)(η¹SC(S)NMe₂)(CO)₂] (R = H, Me) induced by oxidation, Organometallics, 1990, 9, 630–640.
74. C. T. Supuran, Carbonic anhydrases: catalytic and inhibition mechanisms, distribution and physiological roles, Carbonic Anhydrase, CRC Press, 2004, 1, pp. 1–23.
75. V. S. Haritos and G. Dojchinov, Carbonic anhydrase metabolism is a key factor in the toxicity of CO₂ and COS but not CS₂ toward the flour beetle Tribolium castaneum [Coleoptera: Tenebrionidae], Comp. Biochem. Physiol., Part C: Toxicol. Pharmacol., 2005, 140, 139–147.
76. S. Sinnecker, M. Bräuer, W. Koch and E. Anders, CS₂ Fixation by carbonic anhydrase model systems: A new substrate in the catalytic cycle, Inorg. Chem., 2001, 40, 1006–1013.
77. S. Schenk, J. Kesselmeier and E. Anders, How does the exchange of one oxygen atom with sulfur affect the catalytic cycle of carbonic anhydrase?, Chem. – Eur. J., 2004, 10, 3091–3105.
78. J. Notni, S. Schenk, G. Protoschill-Krebs, J. Kesselmeier and E. Anders, The missing link in COS metabolism: a model study on the reactivation of carbonic anhydrase from its hydrosulfide analogue, ChemBioChem, 2007, 8, 530–536.
79. F. Carta, A. Akdemir, A. Scozzafava, E. Masini and C. T. Supuran, Xanthates and trithiocarbonates strongly inhibit carbonic anhydrases and show antiglaucoma effects in vivo, J. Med. Chem., 2013, 56, 4691–4700.
80. M. J. Smeulders, T. R. M. Barends, A. Pol, A. Scherer, M. H. Zandvoort, A. Udvarhelyi, A. F. Khadem, A. Menzel, J. Hermans, R. L. Shoeman, H. J. C. T. Wessels, L. P. van den Heuvel, L. Russ, I. Schlichting, M. S. M. Jetten and H. J. M. Op den Camp, Evolution of a new enzyme for carbon disulphide conversion by an acidothermophilic archaeon, Nature, 2011, 478, 412–416.
81. S. Alcántara, I. Estrada, M. S. Vásquez and S. Revah, Carbon disulfide oxidation by a microbial consortium from a trickling filter, Biotechnol. Lett., 1999, 21, 815–819.
82. N. A. Smith and D. P. Kelly, Oxidation of carbon disulfide as the sole source of energy for the autotrophic growth of Thiobacillus thioparus strain TK-m, J. Gen. Microbiol., 1988, 134, 3041–3048.
83. R. Warington, IV. On nitrification, J. Chem. Soc., Trans., 1878, 33, 44–51.
84. L. C. Seefeldt, M. E. Rasche and S. A. Ensign, Carbonyl sulfide and carbon dioxide as new substrates, and carbon disulfide as a new inhibitor of nitrogenase, Biochemistry, 1995, 34, 5382–5389.
85. M. J. Smeulders, A. Pol, H. Venselaar, T. R. M. Barends, J. Hermans, M. S. M. Jetten and H. J. M. Op den Camp, Bacterial CS₂ hydrolases from Acidithiobacillus thiooxidans strains are homologous to the archaeal catenane CS₂ hydrolase, J. Bacteriol., 2013, 195, 4046–4056.
86. M. R. Hyman, C. Y. Kim and D. J. Arp, Inhibition of ammonia monooxygenase in Nitrosomonas europaea by carbon disulfide, J. Bacteriol., 1990, 172, 4775–4782.
87. M. Kumar, W. P. Lu and S. W. Ragsdale, Binding of carbon disulfide to the site of acetyl-CoA synthesis by the nickel-iron-sulfur protein, carbon monoxide dehydrogenase, from Clostridium thermoaceticum, Biochemistry, 1994, 33, 9769–9777.
88. S. W. Ragsdale and H. G. Wood, Acetate biosynthesis by acetogenic bacteria. Evidence that carbon monoxide dehydrogenase is the condensing enzyme that catalyzes the final steps of the synthesis, J. Biol. Chem., 1985, 260, 3970–3977.
89. R. van Doorn, P. T. Henderson, M. Vanhoorne, P. G. Vertin and C. P. M. J. M. Leijdekkers, Determination of thio compounds in urine of workers exposed to carbon disulfide, Arch. Environ. Health Int. J., 1981, 36, 289–297.
90. M. Silva, A review of developmental and reproductive toxicity of CS₂ and H₂S generated by the pesticide sodium tetrathiocarbonate, Birth Defects Res., Part B, 2013, 98, 119–138.
91. M. J. McKenna and V. DiStefano, Carbon disulfide. I. The metabolism of inhaled carbon disulfide in the rat, J. Pharmacol. Exp. Ther., 1977, 202, 245–252.
92. H. Kivistö, E. Elovaara, V. Riihimäki and A. Aitio, Effect of cytochrome P450 isozyme induction and glutathione depletion on the metabolism of CS₂ to TTCA in rats, Arch. Toxicol., 1995, 69, 185–190.
93. C. V. Smith, D. P. Jones, T. M. Guenthner, L. H. Lash and B. H. Lauterburg, Compartmentation of glutathione: Implications for the study of toxicity and disease, Toxicol. Appl. Pharmacol., 1996, 140, 1–12.
94. D. P. Jones, J. L. Carlson, P. S. Samiec, P. Sternberg Jr., V. C. Mody Jr., R. L. Reed and L. A. S. Brown, Glutathione measurement in human plasma; evaluation of sample collection, storage and derivatization conditions for analysis of dansyl derivatives by HPLC, Clin. Chim. Acta, 1998, 275, 175–184.
95. R. Rossi, D. Giustarini, A. Milzani and I. Dalle-Donne, Cysteinylation and homocysteinylation of plasma protein thiols during ageing of healthy human beings, J. Cell. Mol. Med., 2009, 9B, 3131–3140.
96. S. S. Tang and G. G. Chang, Kinetic characterization of the endogenous glutathione transferase activity of octopus lens S-crystallin, J. Biochem., 1996, 119, 1182–1188.
97. N. Suwandaratne, J. Hu, K. Siriwardana, M. Gadogbe and D. Zhang, Evaluation of thiol raman activities and pK_a values using internally referenced raman-based pH titration, Anal. Chem., 2016, 88, 3624–3631.
98. F. Q. Schafer and G. R. Buettner, Redox environment of the cell as viewed through the redox state of the glutathione disulfide/glutathione couple, Free Radical Biol. Med., 2001, 30, 1191–1212.
99. A. K. Tummanapelli and S. Vasudevan, Ab initio MD simulations of the Brønsted acidity of glutathione in aqueous solutions: predicting pK_a shifts of the cysteine residue, J. Phys. Chem. B, 2015, 119, 15353–15358.
100. S. G. Tajc, B. S. Tolbert, R. Basavappa and B. L. Miller, Direct fetermination of thiol pK_a by isothermal titration microcalorimetry, J. Am. Chem. Soc., 2004, 126, 10508–10509.
101. Oxidative Stress and Redox Regulation, ed. U. Jakob and D. Reichmann, Springer, Netherlands, 2013.
102. J. S. Bus, The relationship of carbon disulfide metabolism to development of toxicity, Neurotoxicology, 1985, 6, 73–80.
103. D. J. Johnson, D. G. Graham, V. Amarnath, K. Amarnath and W. M. Valentine, The measurement of 2-thiothiazolidine-4-carboxylic acid as an index of the in vivo release of CS₂ by dithiocarbamates, Chem. Res. Toxicol., 1996, 9, 910–916.
104. A. Srivastava, S. K. Singh and A. Gupta, Determination of organotrithiocarbonates with o-diacetoxyiodobenzoate and N-chlorosuccinimide in aqueous and non-aqueous media, Analyst, 1990, 115, 421–423.
105. J. E. Drake and J. Yang, Preparation and characterization of trimethylgermanium−sulfur compounds derived from 2-dithiocarbamoyl-3-dithiocarbonylthiopropionate. Crystal Structures of [NH₄]₃[S₂CSCH₂CH(NHCS₂)CO₂] and Me₃GeO₂C(−CHNHC(S)SCH₂−), Inorg. Chem., 1998, 37, 2968–2974.
106. L. Magos and J. A. E. Jarvis, Effects of diethyldithiocarbamate and carbon disulphide on brain tyrosine, J. Pharm. Pharmacol., 1970, 22, 936–938.
107. M. J. McKenna and V. DiStefano, Carbon disulfide. II. A proposed mechanism for the action of carbon disulfide on dopamine β-hydroxylase, J. Pharmacol. Exp. Ther., 1977, 202, 253–266.
108. R. J. Rubin, Role of metabolic activation in the renal toxicity of carbon disulfide (CS₂), Toxicol. Lett., 1990, 53, 211–213.
109. R. F. M. Herber and H. Poppe, A new method for the estimation of exposure to carbon disulphide, J. Chromatogr. A, 1976, 118, 23–34.
110. R. E. Heikkila, F. S. Cabbat and G. Cohen, In vivo inhibition of superoxide dismutase in mice by diethyldithiocarbamate, J. Biol. Chem., 1976, 251, 2182–2185.
111. M. Wiedau-Pazos, J. J. Goto, S. Rabizadeh, E. B. Gralla, J. A. Roe, M. K. Lee, J. S. Valentine and D. E. Bredesen, Altered reactivity of superoxide dismutase in familial amyotrophic lateral sclerosis, Science, 1996, 271, 515–518.
112. M. Bozdag, F. Carta, D. Vullo, A. Akdemir, S. Isik, C. Lanzi, A. Scozzafava, E. Masini and C. T. Supuran, Synthesis of a new series of dithiocarbamates with effective human carbonic anhydrase inhibitory activity and antiglaucoma action, Bioorg. Med. Chem., 2015, 23, 2368–2376.
113. L. A. Shinobu, S. G. Jones and M. M. Jones, Mobilization of aged cadmium deposits by dithiocarbamates, Arch. Toxicol., 1983, 54, 235–242.
114. F. W. Sunderman, Chelation therapy in nickel poisoning, Ann. Clin. Lab. Sci., 1981, 11, 1–8.
115. E. C. Reisinger, P. Kern, M. Dietrich, M. Ernst, H. D. Flad, P. Bock and German Dtc Study Group, Inhibition of HIV progression by dithiocarb, Lancet, 1990, 335, 679–682.
116. E. M. Hersh, G. Brewton, D. Abrams, J. Bartlett, J. Galpin, P. Gill, R. Gorter, M. Gottlieb, J. J. Jonikas, S. Landesman, A. Levine, A. Marcel, E. A. Petersen, M. Whiteside, J. Zahradnik, C. Negron, F. Boultitie, J. Caraux, J. M. Dupuy and R. Salmi, Ditiocarb sodium (diethyldithiocarbamate) therapy in patients with symptomatic HIV infection and AIDS: a randomized, double-blind, placebo-controlled, multicenter study, JAMA, J. Am. Med. Assoc., 1991, 265, 1538–1544.
117. V. Bala, S. Jangir, D. Mandalapu, S. Gupta, Y. S. Chhonker, N. Lal, B. Kushwaha, H. Chandasana, S. Krishna, K. Rawat, J. P. Maikhuri, R. S. Bhatta, M. I. Siddiqi, R. Tripathi, G. Gupta and V. L. Sharma, Dithiocarbamate–thiourea hybrids useful as vaginal microbicides also show reverse transcriptase inhibition: design, synthesis, docking and pharmacokinetic studies, Bioorg. Med. Chem. Lett., 2015, 25, 881–886.
118. H. Mansour, M. Levacher, M. A. Gougerot-Pocidalo, B. Rouveix and J. J. Pocidalo, Diethyldithiocarbamate provides partial protection against pulmonary and lymphoid oxygen toxicity, J. Pharmacol. Exp. Ther., 1986, 236, 476–480.
119. S. Orrenius, C. S. I. Nobel, D. J. van den Dobbelsteen, M. J. Burkitt and A. F. G. Slater, Dithiocarbamates and the redox regulation of cell death, Biochem. Soc. Trans., 1996, 24, 1032–1038.
120. G. Vettorazzi, W. F. Almeida, G. J. Burin, R. B. Jaeger, F. R. Puga, A. F. Rahde, F. G. Reyes and S. Schvartsman, International safety assessment of pesticides: dithiocarbamate pesticides, ETU, and PTU – A review and update, Teratog., Carcinog., Mutagen., 1995, 15, 313–337.
121. C. Mathieu, R. Duval, X. Xu, F. Rodrigues-Lima and J. M. Dupret, Effects of pesticide chemicals on the activity of metabolic enzymes: focus on thiocarbamates, Expert Opin. Drug Metab. Toxicol., 2014, 11, 1–14.
122. A. Moretto and C. Colosio, Biochemical and toxicological evidence of neurological effects of pesticides: the example of Parkinson's disease, NeuroToxicology, 2011, 32, 383–391.
123. T. P. Brown, P. C. Rumsby, A. C. Capleton, L. Rushton and L. S. Levy, Pesticides and parkinson's disease-is there a link?, Environ. Health Perspect., 2006, 114, 156–164.
124. Y. Zhou, F. S. Shie, P. Piccardo, T. J. Montine and J. Zhang, Proteasomal inhibition induced by manganese ethylene-bis-dithiocarbamate: relevance to Parkinson's disease, Neuroscience, 2004, 128, 281–291.
125. N. Edington and J. M. Howell, Changes in the nervous system of rabbits following the administration of sodium diethyldithiocarbamate, Nature, 1966, 210, 1060–1062.
126. A. R. Rasul and J. M. Howell, Further observations on the response of the peripheral and central nervous system of the rabbit to sodium diethyldithiocarbamate, Acta Neuropathol., 1973, 24, 161–173.
127. G. B. Frisoni and V. Monda, Disulfiram neuropathy: a review (1971–1988) and report of a case, Alcohol Alcohol., 1989, 24, 429–437.
128. B. Mokri, A. Ohnishi and P. J. Dyck, Disulfiram neuropathy, Neurology, 1981, 31, 730.
129. W. M. Valentine, V. Amarnath, K. Amarnath, F. Rimmele and D. G. Graham, Carbon disulfide mediated protein crosslinking by N,N-diethyldithiocarbamate, Chem. Res. Toxicol., 1995, 8, 96–102.
130. E. G. Tonkin, J. C. L. Erve and W. M. Valentine, Disulfiram produces a non-carbon disulfide-dependent schwannopathy in the rat, J. Neuropathol. Exp. Neurol., 2000, 59, 786–797.
131. D. I. Eneanya, J. R. Bianchine, D. O. Duran and B. D. Andresen, The actions and metabolic fate of disulfiram, Annu. Rev. Pharmacol. Toxicol., 1981, 21, 575–596.
132. B. Cvek and Z. Dvorak, Targeting of nuclear factor-κB and proteasome by dithiocarbamate complexes with metals, Curr. Pharm. Des., 2007, 13, 3155–3167.
133. P. I. Nagy, G. Alagona, C. Ghio and K. Takács-Novák, Theoretical conformational analysis for neurotransmitters in the gas phase and in aqueous solution. Norepinephrine, J. Am. Chem. Soc., 2003, 125, 2770–2785.
134. R. Brisch, A. Saniotis, R. Wolf, H. Bielau, H. G. Bernstein, J. Steiner, B. Bogerts, K. Braun, Z. Jankowski, J. Kumaratilake, M. Henneberg and T. Gos, The role of dopamine in schizophrenia from a neurobiological and evolutionary perspective: old fashioned, but still in vogue, Front. Mol. Psychiatry, 2014, 5 DOI:10.3389/fpsyt.2014.00047.
135. F. L. O'Connor, The role of serotonin and dopamine in schizophrenia, J. Am. Psychiatr. Nurses Assoc., 1998, 4, S30–S34.
136. K. A. Veverka, K. L. Johnson, D. C. Mays, J. J. Lipsky and S. Naylor, Inhibition of aldehyde dehydrogenase by disulfiram and its metabolite methyl diethylthiocarbamoyl-sulfoxide, Biochem. Pharmacol., 1997, 53, 511–518.
137. J. P. Schroeder, D. A. Cooper, J. R. Schank, M. A. Lyle, M. Gaval-Cruz, Y. E. Ogbonmwan, N. Pozdeyev, K. G. Freeman, P. M. Iuvone, G. L. Edwards, P. V. Holmes and D. Weinshenker, Disulfiram attenuates drug-primed reinstatement of cocaine seeking via inhibition of dopamine β-hydroxylase, Neuropsychopharmacology, 2010, 35, 2440–2449.
138. J. J. Lipsky, M. L. Shen and S. Naylor, In vivo inhibition of aldehyde dehydrogenase by disulfiram, Chem.-Biol. Interact., 2001, 130, 93–102.
139. K. S. Barth and R. J. Malcolm, Disulfiram: an old therapeutic with new applications, CNS Neurol. Disord.: Drug Targets, 2010, 9, 5–12.
140. L. Z. Lin and J. Lin, Antabuse (disulfiram) as an affordable and promising anticancer drug, Int. J. Cancer, 2011, 129, 1285–1286.
141. J. C. L. Erve, V. Amarnath, D. G. Graham, R. C. Sills, A. L. Morgan and W. M. Valentine, Carbon disulfide and N,N-diethyldithiocarbamate generate thiourea cross-links on erythrocyte spectrin in vivo, Chem. Res. Toxicol., 1998, 11, 544–549.
142. R. C. Sills, W. M. Valentine, V. Moser, D. G. Graham and D. L. Morgan, Characterization of carbon disulfide neurotoxicity in C57BL6 mice: behavioral, morphologic, and molecular effects, Toxicol. Pathol., 2000, 28, 142–148.
143. D. G. Isom, C. A. Castañeda, B. R. Cannon and E. B. García-Moreno, Large shifts in pK_a values of lysine residues buried inside a protein, Proc. Natl. Acad. Sci. U. S. A., 2011, 108, 5260–5265.
144. A. P. DeCaprio, D. C. Spink, X. Chen, J. H. Fowke, M. Zhu and S. Bank, Characterization of isothiocyanates, thioureas, and other lysine adduction products in carbon disulfide-treated peptides and protein, Chem. Res. Toxicol., 1992, 5, 496–504.
145. R. Schreck, B. Meier, D. N. Männel, W. Dröge and P. A. Baeuerle, Dithiocarbamates as potent inhibitors of nuclear factor κB activation in intact cells, J. Exp. Med., 1992, 175, 1181–1194.
146. M. Karin and F. R. Greten, NF-κB: linking inflammation and immunity to cancer development and progression, Nat. Rev. Immunol., 2005, 5, 749–759.
147. X. Dolcet, D. Llobet, J. Pallares and X. Matias-Guiu, NF-κB in development and progression of human cancer, Virchows Arch., 2005, 446, 475–482.
148. J. Dutta, Y. Fan, N. Gupta, G. Fan and C. Gelinas, Current insights into the regulation of programmed cell death by NF-κB, Oncogene, 2006, 25, 6800–6816.
149. S. Beinke and S. C. Ley, Functions of NF-kB1 and NF-kB2 in immune cell biology, Biochem. J., 2004, 382, 393–409.
150. M. S. Hayden, A. P. West and S. Ghosh, NF-κB and the immune response, Oncogene, 2006, 25, 6758–6780.
151. D. S. Basseres and A. S. Baldwin, Nuclear factor-κB and inhibitor of κB kinase pathways in oncogenic initiation and progression, Oncogene, 2006, 25, 6817–6830.
152. T. D. Gilmore and M. Herscovitch, Inhibitors of NF-κB signaling: 785 and counting, Oncogene, 2006, 25, 6887–6899.
153. G. Bonizzi and M. Karin, The two NF-κB activation pathways and their role in innate and adaptive immunity, Trends Immunol., 2004, 25, 280–288.
154. H. L. Pahl, Activators and target genes of Rel/Nf-kB transcription factors, Oncogene, 1999, 18, 6853–6866.
155. C. Morais, B. Pat, G. Gobe, D. W. Johnson and H. Healy, Pyrrolidine dithiocarbamate exerts anti-proliferative and pro-apoptotic effects in renal cell carcinoma cell lines, Nephrol., Dial., Transplant., 2006, 21, 3377–3388.
156. S. Miyamoto and I. M. Verma, REL/NF-κB/IκB Story, Adv. Cancer Res., 1995, 66, 255–292.
157. M. J. May and S. Ghosh, Rel/NF-κB and IκB proteins: an overview, Semin. Cancer Biol., 1997, 8, 63–73.
158. P. A. Baeuerle, IκB–NF-κB structures: at the interface of inflammation control, Cell, 1998, 95, 729–731.
159. C. Scheidereit, IκB kinase complexes: gateways to NF-κB activation and transcription, Oncogene, 2006, 25, 6685–6705.
160. M. Hayakawa, H. Miyashita, I. Sakamoto, M. Kitagawa, H. Tanaka, H. Yasuda, M. Karin and K. Kikugawa, Evidence that reactive oxygen species do not mediate NF-κB activation, EMBO J., 2003, 22, 3356–3366.
161. K. H. Ha, M. S. Byun, J. Choi, J. Jeong, K. J. Lee and D. M. Jue, N-Tosyl-l-phenylalanine chloromethyl ketone inhibits NF-κB activation by blocking specific cysteine residues of IκB Kinase β and p65/RelA, Biochemistry, 2009, 48, 7271–7278.
162. H. Sebban, S. Yamaoka and G. Courtois, Posttranslational modifications of NEMO and its partners in NF-κB signaling, Trends Cell Biol., 2006, 16, 569–577.
163. N. L. Reynaert, A. van der Vliet, A. S. Guala, T. McGovern, M. Hristova, C. Pantano, N. H. Heintz, J. Heim, Y. S. Ho, D. E. Matthews, E. F. M. Wouters and Y. M. W. Janssen-Heininger, Dynamic redox control of NF-κB through glutaredoxin-regulated S-glutathionylation of inhibitory κB kinase β, Proc. Natl. Acad. Sci. U. S. A., 2006, 103, 13086–13091.
164. M. S. Byun, J. Choi and D. M. Jue, Cysteine-179 of IκB kinase β plays a critical role in enzyme activation by promoting phosphorylation of activation loop serines, Exp. Mol. Med., 2006, 38, 546–552.
165. S. Liu, Y. R. Misquitta, A. Olland, M. A. Johnson, K. S. Kelleher, R. Kriz, L. L. Lin, M. Stahl and L. Mosyak, Crystal structure of a human IκB kinase β asymmetric dimer, J. Biol. Chem., 2013, 288, 22758–22767.
166. K. I. Jeon, M. S. Byun and D. M. Jue, Gold compound auranofin inhibits IκB kinase (IKK) by modifying Cys-179 of IKKβ subunit, Exp. Mol. Med., 2003, 35, 61–66.
167. K. Heyninck, M. Lahtela-Kakkonen, P. Van der Veken, G. Haegeman and W. Vanden Berghe, Withaferin A inhibits NF-κB activation by targeting cysteine 179 in IKKβ, Biochem. Pharmacol., 2014, 91, 501–509.
168. C. S. Lai, Methods for the controlled delivery of carbon disulfide for the treatment of inflammatory conditions, WO Pat. App., WO1999040907 A1, US, 1999.
169. F. De Matteis, Covalent binding of sulfur to microsomes and loss of cytochrome P-450 during the oxidative desulfuration of several chemicals, Mol. Pharmacol., 1974, 10, 849–854.
170. F. De Matteis and A. A. Seawright, Liver toxicity of carbon disulfide. Possible significance of its metabolism by oxidative desulphuration, Panminerva Med., 1976, 18, 364–366.
171. C. J. Decker, D. R. Doerge and J. R. Cashman, Metabolism of benzimidazoline-2-thiones by rat hepatic microsomes and hog liver flavin-containing monooxygenase, Chem. Res. Toxicol., 1992, 5, 726–733.
172. C. P. Chengelis and R. A. Neal, Hepatic carbonyl sulfide metabolism, Biochem. Biophys. Res. Commun., 1979, 90, 993–999.
173. R. A. Neal, Microsomal metabolism of thiono-sulfur compounds: mechanisms and toxicological significance, Rev. Biochem. Toxicol., 1980, 2, 131–171.
174. J. Halpert and R. A. Neal, Inactivation of rat liver cytochrome P-450 by the suicide substrates parathion and chloramphenicol, Drug Metab. Rev., 1981, 12, 239–259.
175. Q. Li and J. R. Lancaster Jr., Chemical foundations of hydrogen sulfide biology, Nitric Oxide, 2013, 35, 21–34.
176. K. G. Reddie and K. S. Carroll, Expanding the functional diversity of proteins through cysteine oxidation, Curr. Opin. Chem. Biol., 2008, 12, 746–754.
177. T. Ida, T. Sawa, H. Ihara, Y. Tsuchiya, Y. Watanabe, Y. Kumagai, M. Suematsu, H. Motohashi, S. Fujii, T. Matsunaga, M. Yamamoto, K. Ono, N. O. Devarie-Baez, M. Xian, J. M. Fukuto and T. Akaike, Reactive cysteine persulfides and S-polythiolation regulate oxidative stress and redox signaling, Proc. Natl. Acad. Sci., U. S. A., 2014, 111, 7606–7611.
178. A. K. Mustafa, M. M. Gadalla and S. H. Snyder, Signaling by gasotransmitters, Sci. Signaling, 2009, 2(68) DOI:10.1126/scisignal.268re2.
179. K. Ono, T. Akaike, T. Sawa, Y. Kumagai, D. A. Wink, D. J. Tantillo, A. J. Hobbs, P. Nagy, M. Xian, J. Lin and J. M. Fukuto, The redox chemistry and chemical biology of H₂S, hydropersulfides and derived species: implications of their possible biological activity and utility, Free Radical Biol. Med., 2014, 77, 82–94.
180. S. S. Saund, V. Sosa, S. Henriquez, Q. N. N. Nguyen, C. L. Bianco, S. Soeda, R. Millikin, C. White, H. Le, D. J. Tantillo, Y. Kumagai, T. Akaike, J. Lin and J. M. Fukuto, The chemical biology of hydropersulfides (RSSH): chemical stability, reactivity and redox roles, Arch. Biochem. Biophys., 2015, 588, 15–24.
181. R. J. Hondal, S. M. Marino and V. N. Gladyshev, Selenocysteine in thiol/disulfide-like exchange reactions, Antioxid. Redox Signalling, 2013, 18, 1675–1689.
182. J. K. Wrobel, R. Power and M. Toboek, Biological activity of selenium: revisited, IUBMB Life, 2016, 68, 97–105.
183. Y. M. Go, J. D. Chandler and D. P. Jones, The cysteine proteome, Free Radical Biol. Med., 2015, 84, 227–245.
184. B. Buszewski, A. Ulanowska, T. Ligor, M. Jackowski, E. Kłodzińska and J. Szeliga, Identification of volatile organic compounds secreted from cancer tissues and bacterial cultures, J. Chromatogr. B: Anal. Technol. Biomed. Life Sci., 2008, 868, 88–94.
185. W. Miekisch, J. K. Schubert and G. F. Noeldge-Schomburg, Diagnostic potential of breath analysis – focus on volatile organic compounds, Clin. Chim. Acta, 2004, 347, 25–39.
186. M. Phillips, Detection of carbon disulfide in breath and air: a possible new risk factor for coronary artery disease, Int. Arch. Occup. Environ. Health, 1992, 64, 119–123.
187. M. Phillips, M. Sabas and J. Greenberg, Increased pentane and carbon disulfide in the breath of patients with schizophrenia, J. Clin. Pathol., 1993, 46, 861–864.
188. B. Vitali, M. Ndagijimana, F. Cruciani, P. Carnevali, M. Candela, M. E. Guerzoni and P. Brigidi, Impact of a symbiotic food on the gut microbial ecology and metabolic profiles, BMC Microbiol., 2010, 10(4) DOI:10.1186/1471-2180-10-4.
189. B. Vitali, M. Ndagijimana, S. Maccaferri, E. Biagi, M. E. Guerzoni and P. Brigidi, An in vitro evaluation of the effect of probiotics and prebiotics on the metabolic profile of human microbiota, Anaerobe, 2012, 18, 386–391.
190. S. Studer, J. Orens, I. Rosas, J. Krishnan, K. Cope, S. Yang, J. Conte, P. Becker and T. Risby, Patterns and significance of exhaled-breath biomarkers in lung transplant recipients with acute allograft rejection, J. Heart Lung Transplant., 2001, 20, 1158–1166.
191. B. G. Galef Jr., J. R. Mason, G. Preti and N. J. Bean, Carbon disulfide: a semiochemical mediating socially-induced diet choice in rats, Physiol. Behav., 1988, 42, 119–124.
192. S. D. Munger, T. Leinders-Zufall, L. M. McDougall, R. E. Cockerham, A. Schmid, P. Wandernoth, G. Wennemuth, M. Biel, F. Zufall and K. R. Kelliher, An Olfactory Subsystem that Detects Carbon Disulfide and Mediates Food-Related Social Learning, Curr. Biol., 2010, 20, 1438–1444.
193. H. Arakawa, K. R. Kelliher, F. Zufall and S. D. Munger, The receptor guanylyl cyclase type D (GC-D) ligand uroguanylin promotes the acquisition of food preferences in mice, Chem. Senses, 2013, 38, 391–397.
194. World Health Organization, Carbon disulfide, Air Quality Guidelines, WHO Regional Office for Europe, Copenhagen, Denmark, 2nd edn, 2000, ch. 5.4.
195. T. Göen, A. Schramm, T. Baumeister, W. Uter and H. Drexler, Current and historical individual data about exposure of workers in the rayon industry to carbon disulfide and their validity in calculating the cumulative dose, Int. Arch. Occup. Environ. Health, 2013, 87, 675–683.
196. A. Schramm, W. Uter, M. Brandt, T. Göen, M. Köhrmann, T. Baumeister and H. Drexler, Increased intima-media thickness in rayon workers after long-term exposure to carbon disulfide, Int. Arch. Occup. Environ. Health, 2015, 89, 513–519.
197. A. Eben and F. P. Freudlsperger, 2-Thioxothiazolidine-4-carboxylic acid (TTCA) [Biomonitoring Methods, 1994], The MAK-Collection for Occupational Health and Safety, Wiley-VCH Verlag GmbH & Co. KGaA, 2002 DOI:10.1002/3527600418.
198. V. Riihimäki, H. Kivistö, K. Peltonen, E. Helpiö and A. Aitio, Assessment of exposure to carbon disulfide in viscose production workers from urinary 2-thiothiazolidine-4-carboxylic acid determinations, Am. J. Ind. Med., 1992, 22, 85–97.
199. L. Campbell, A. H. Jones and H. K. Wilson, Evaluation of occupational exposure to carbon disulphide by blood, exhaled air, and urine analysis, Am. J. Ind. Med., 1985, 8, 143–153.
200. J. Rosier, M. Vanhoorne, R. Grosjean, E. Van De Walle, G. Billemont and C. Van Peteghem, , Preliminary evaluation of urinary 2-thio-thiazolidine-4-carboxylic-acid (TTCA) levels as a test for exposure to carbon disulfide, Int. Arch. Occup. Environ. Health, 1982, 51, 159–167.
201. R. van Doorn, L. P. C. Delbressine, C. M. Leijdekkers, P. G. Vertin and P. T. Henderson, Identification and determination of 2-thiothiazolidine-4-carboxylic acid in urine of workers exposed to carbon disulfide, Arch. Toxicol., 1981, 47, 51–58.
202. J. Roh, C. N. Kim, N. G. Lim, J. H. Chang and Y. B. Cho, Simultaneous analysis of urinary 2-thiothiazolidine-4-carboxylic acid and thiocarbamide as a biological exposure index for carbon disulfide exposure, Yonsei Med. J., 1999, 40, 265–272.
203. L. A. Ridnour, D. D. Thomas, C. Switzer, W. Flores-Santana, J. S. Isenberg, S. Ambs, D. D. Roberts and D. A. Wink, Molecular mechanisms for discrete nitric oxide levels in cancer, Nitric Oxide, 2008, 19, 73–76.
204. P. Simon, T. Nicot and M. Dieudonné, Dietary habits, a non-negligible source of 2-thiothiazolidine-4-carboxylic acid and possible overestimation of carbon disulfide exposure, Int. Arch. Occup. Environ. Health, 1994, 66, 85–90.
205. Y. Kikuchi, T. Uemura, T. Yamuchi, T. Takebayashi, Y. Nishiwaki, K. Yamada, H. Sakurai and K. Omae, Urinary excretion of TTCA after intake of brassica vegetables, J. Occup. Health, 2002, 44, 151–155.
206. International Agency for Research on Cancer, Cruciferous vegetables, isothiocyanates and Indoles, IARC Press, Lyon, France, vol. 9, 2004.
207. P. Riso, C. Del Bo' and S. Vendrame, Preventive effects of broccoli bioactives: role on oxidative stress and cancer risk, in Cancer: Oxidative Stress and Dietary Antioxidants, ed. V. Preedy, Elsevier Inc., Academic Press, San Diego, 2014, ch. 11, pp. 115–126.
208. L. K. Keefer, R. W. Nims, K. M. Davies and D. A. Wink, ‘NONOates’ (1-substituted diazen-1-ium-1,2-diolates) as nitric oxide donors: convenient nitric oxide dosage forms, Methods Enzymol., 1996, 268, 281–293.
209. A. E. Pierri, D. A. Muizzi, A. D. Ostrowski and P. C. Ford, Photo-controlled release of NO and CO with inorganic and organometallic complexes, Struct. Bonding, 2014, 165, 1–45.

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