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Biomolecules 2014, 4(2), 419-434; doi:10.3390/biom4020419
Published: 17 April 2014
Abstract: Cysteine residues are known to perform essential functions within proteins, including binding to various metal ions. In particular, cysteine residues can display high affinity toward zinc ions (Zn2+), and these resulting Zn2+-cysteine complexes are critical mediators of protein structure, catalysis and regulation. Recent advances in both experimental and theoretical platforms have accelerated the identification and functional characterization of Zn2+-bound cysteines. Zn2+-cysteine complexes have been observed across diverse protein classes and are known to facilitate a variety of cellular processes. Here, we highlight the structural characteristics and diverse functional roles of Zn2+-cysteine complexes in proteins and describe structural, computational and chemical proteomic technologies that have enabled the global discovery of novel Zn2+-binding cysteines.
When considering biologically relevant transition metals, zinc is the second most abundant found within cells, behind only iron. Zinc ions (Zn2+) are known to facilitate diverse protein functions that are essential for life. Common Zn2+ ligands found within proteins include cysteine (S), histidine (N), aspartate (O), and glutamate (O) residues. The ionization state of the thiol group of cysteine governs its ability to bind metals, including Zn2+. The pKa of the thiol group of cysteine is typically close to physiological pH (7.4) ; therefore, the ionization state of cysteine is highly sensitive to small changes within the local protein environment . Thus, the affinity of cysteine for Zn2+ varies accordingly for each individual cysteine within a protein scaffold. These resulting complexes contribute to protein structure, catalysis, and regulation (Figure 1) [3,4]. Zn2+-containing protein structural motifs are best highlighted by the well-characterized zinc fingers, first discovered over 25 years ago [5,6,7]. Although less common, Zn2+-cysteine complexes also catalyze enzymatic transformations in diverse classes such as oxidoreductases, transferases, and hydrolases. More recently, the potential for Zn2+ to modulate protein activities has been established. These regulatory complexes proceed through distinct mechanisms, such as Zn2+-inhibition, redox-switches, and stabilization of protein interfaces [8,9,10]. Lastly, the cysteine-rich metallothioneins tightly regulate cellular Zn2+ levels by storing and properly redistributing Zn2+ throughout the cell . Due to these diverse functional roles of Zn2+-cysteine complexes, the development of both experimental and theoretical approaches has been paramount in the identification and characterization of Zn2+-binding cysteines. Here we summarize key examples of functional Zn2+-cysteine complexes, and discuss recent advances in computational and proteomic technologies to study these complexes.
2. Structural Zn2+-Cysteine Complexes: Zinc Fingers
Because zinc is a d10 transition metal, it exclusively forms a Zn2+ ion and typically lacks redox activity within cells. Zn2+ typically is found to assemble coordination complexes with four ligands in a tetrahedral geometry. Recent studies estimate the human proteome consists of approximately 3000 Zn2+-proteins . Of the potential Zn2+ ligands within proteins, the sulfur atom of cysteine transfers the most charge over to the Zn2+. As cysteine occupies more ligand sites, it often quenches the ability of the Zn2+ to act as a lewis acid, rendering these complexes relatively inert . As a result, Zn2+-cysteine complexes traditionally perform structural roles within proteins. The most abundant class of structural Zn2+-cysteine complexes is the zinc finger, and these have been extensively classified . Zinc fingers are characteristically comprised of Cys4 or Cys2His2 coordination environments . The classical Cys2His2 zinc finger chelates a single Zn2+ within an α-helix and antiparallel β-sheet (Figure 2 inset) . Zinc finger domains are typically found in clusters of four or more within a single protein, and often structurally stabilize the protein for interaction with other proteins and biomolecules, such as DNA and RNA.
Although the functional roles of most zinc finger proteins are poorly understood, most annotated proteins act as transcription activators or suppressors . A single zinc finger possesses four amino acids at the −1, 2, 3, and 6 positions of the α-helix that participate in hydrogen-bond interactions with 3–4 nucleic acids within the major groove of DNA (Figure 2) . Different sequences at these four positions preferentially bind to distinct nucleic acid sequences with high affinity and selectivity . Consequently, this motif has been exploited in the development of zinc finger endonucleases for genetic engineering. By conjugating specific arrays of zinc fingers to a promiscuous FokI endonuclease, DNA can be cut at an indicated sequence to disrupt, add, or correct the gene of interest . The development of a conserved linker sequence was vital to the construction of polymeric zinc-finger endonucleases, requiring DNA sequences of up to 18 bp for recognition . This advance provided enough specificity to target single genes within the human genome [19,20]. The expansion of synthetic zinc finger endonucleases has extended genetic engineering across diverse gene families .
3. Catalytic Zn2+-Cysteine Complexes
Beyond structural roles, cysteines bind Zn2+ to directly facilitate enzymatic transformations. Cysteines are less common ligands in catalytic Zn2+ complexes due to the steric bulk of sulfur and greater charge transfer compared to histidine and water ligands . However, catalytic Zn2+-cysteine complexes have been observed across diverse enzyme classes, such as oxidoreductases, hydrolases, and transferases (Table 1). The exact mechanism varies within each individual enzyme, but typically is comprised of either substrate coordination or activation by Zn2+. Alcohol dehydrogenase enzymes (ADH) were first discovered to require a Zn2+ for catalysis over 50 years ago . These evolutionarily conserved enzymes facilitate the interconversion between alcohols and ketones or aldehydes. Humans possess six distinct classes of ADH enzymes (ADH1-ADH6), each utilizing the Zn2+-dependent catalytic mechanism . The active enzyme is a dimer, with each 40 kD monomer possessing a substrate-binding Zn2+ and an NAD+ cofactor [23,24]. In the case of ADH5, the catalytic Zn2+ is bound to Cys46, His66, Cys174, and the alcohol substrate (Figure 3a) . The bound Zn2+ coordinates the substrate in the correct geometry for the sequential proton transfer to Ser48 followed by hydride transfer to NAD+ (Figure 3b) . Although they do not directly interact with the substrate, these cysteine residues are highly conserved throughout human ADH enzyme classes and are essential for ADH activity .
|Table 1. Representative human proteins containing catalytic Zn2+-cysteine complexes.|
|Protein||Enzyme Class||Function||Mechanism||PDB Structure|
|Alcohol dehydrogenase||Oxidoreductase||Interconverts alcohols to aldehydes and ketones||Zn2+-coordination of substrate ||1MC5 |
|Sorbitol dehydrogenase||Oxidoreductase||Reversible conversion of sorbitol to fructose||Zn2+-activation of nucleophilic water molecule ||1PL7 |
|Cytidine deaminase||Hydrolase||Irreversible hydrolytic deamination of cytidine to uridine||Zn2+-activation of nucleophilic water molecule [28,29]||2KEM |
|GTP cyclohydrolase I||Hydrolase||Converts GTP to dihydroneopterin triphosphate||Zn2+-activation of nucleophilic water molecule ||1FB1 |
|Betain-homocysteine methyltransferase||Transferase||Transfers methyl group from betaine to homocysteine, forming dimethyl glycine and methionine||Zn2+-activation of thiol of homocysteine substrate ||1LT8 |
|Protein farnesyltransferase||Transferase||Post-translational addition of farnesyl to cysteine residues within proteins||Zn2+-activation of thiol on target protein [33,34]||1JCQ |
While ADH acts through a Zn2+-substrate coordination mechanism, farnesyl transferase (FTase) relies on the activation of the substrate thiol by Zn2+ for its activity. FTase is part of the prenyltransferase protein family and catalyzes the post-translational addition of the 15-carbon farnesyl isoprenoid to proteins such as Ras, Rho, and Rab [36,37]. The isoprenoid is attached through a thioether linkage to a cysteine residue within a C-terminal CaaX peptide and is required for proper protein function by mediating membrane association and protein-protein interactions . A Zn2+ is coordinated to Asp297, Cys299, and His362 within the active site of the β subunit of FTase (Figure 3c) . The cysteine residue of the protein substrate coordinates to the Zn2+, displacing either a water or an Asp ligand. The adjacently bound farnesyl diphosphate is now vulnerable to nucleophilic attack by the Zn2+-activated thiol, resulting in the release of inorganic phosphate (PPi) and the farnesylated protein (Figure 3d) [33,34]. It’s important to note that ADH and FTase represent only two possible Zn2+-depended enzymatic mechanisms, and many others have been observed as well. In an alternative mechanism, cytodine deaminase employs a Zn2+ bound by a histidine, two cysteines, and a water molecule to irreversibly deaminate cytidine to uridine . In this case, the water molecule becomes activated and acts as the nucleophile that facilitates the transformation [28,29]. These described catalytic mechanisms refute the common misconception that Zn2+-cysteine complexes are only capable of serving structural roles within proteins.
4. Regulatory Zn2+-Cysteine Complexes
Additionally, cysteine residues also have been observed to bind Zn2+ to modulate protein activities. In these cases, Zn2+-binding must be more transient in nature to allow for interchange between the bound and apo-forms. As a result, these cysteines are often more challenging to identify. Characterized regulatory mechanisms range in complexity, and have been categorized as inhibitory, redox-switches, and protein interface Zn2+-cysteine complexes (Table 2).
|Table 2. Representative human proteins containing regulatory Zn2+-cysteine complexes.|
|Protein||Enzyme Class||Function||Mechanism||PDB Structure|
|Dimethylarginine dimethylaminohydrolase||Hydrolase||Converts N-omega,N-omega-methyl-L-arginine to dimethylamine and L-citrulline||Inhibitory ||2CI7 |
|Ornithine transcarbamoylase||Transferase||Converts carbamoyl phosphate and ornithine to citrulline and phosphate||Inhibitory ||1EP9 |
|Cathepsin S||Protease||Lysosomal cysteine protease||Inhibitory [8,43]||2HH5 |
|Caspase 3||Protease||Cysteine protease||Inhibitory [44,45]||-|
|Caspase 6||Protease||Cysteine protease||Inhibitory ||4FXO |
|Caspase 9||Protease||Cysteine protease||Inhibitory ||1JXQ |
|Aconitase 2||Isomerase||Converts citrate to iso-citrate||Inhibitory ||-|
|Glutathione S-transferase omega||Transferase||Conjugates glutathione to a variety of electrophiles||Inhibitory ||-|
|Betain-homocysteine methyltransferase||Transferase||Transfers methyl group from betaine to homocysteine, forming dimethyl glycine and methionine||Redox-switch ||1LT7, 1LT8 |
|Protein kinase C||Kinase||Phosphorylates serines and threonines||Redox-switch ||3PFQ |
|Nitric oxide synthase||Oxidoreductase||Produces nitric oxide from arginine||Protein interface; Redox-switch ||3NOS |
|Apo2L/TRAIL||Cytokine||Induces signaling pathways to trigger apoptosis||Protein interface ||1DG6 |
4.1. Inhibitory Zn2+-Cysteine Complexes
Cysteine residues have been found to bind Zn2+ as a means of inhibiting enzymatic activity . Inhibition usually occurs by chelation of Zn2+ to the catalytic cysteine residue, but allosteric inhibition attributed to Zn2+-binding at a cysteine distal to the active site has also been described (Table 2).
Dimethylarginine dimethylaminohydrolase (DDAH-1) is a metabolic enzyme responsible for the conversion of dimethylarginine to dimethylamine and citrulline. Dimethylarginine is known to inhibit nitric oxide synthases to mitigate the production of nitric oxide, an important cell signaling molecule . The most well-studied DDAH-1 is from bovine, however, the human homologue retains 94% sequence homology. Zn2+ inhibits DDAH-1 activity with a Ki of 4.2 nM at pH 7.4 . This value is rather high when considering the physiological range of available Zn2+ concentrations and is suggestive of a weaker, more transient binding mode that is indicative of a regulatory role for Zn2+ within DDAH-1. The enzyme functions through a nucleophilic cysteine residue (Cys274) conserved in both the human and bovine proteins . Structural studies reveal a Zn2+ bound to the catalytic Cys274 and His173 within the active site of the enzyme (Figure 4a) . The remaining two ligands are comprised of water molecules stabilized by hydrogen-bonding to adjacent Asp79 and Glu78. DDAH-1 only possesses two Zn2+ ligands instead of the typical three or four, which may contribute to the weaker, more transient Zn2+ binding.
Although most inhibitory Zn2+-cysteine complexes are found to bind directly to the nucleophilic cysteine residue, the potential for allosteric inhibition has been realized in the case of Caspase-9. Caspases are cysteine-dependent aspartate-directed proteases that play a prevalent role in signaling cascades culminating in apoptosis . Zn2+ has been implicated as a strict mediator of apoptosis, where small fluctuations in concentration can strongly dictate cell survival or death . Caspase-9 is an initiator caspase that goes on to cleave caspase-3 and 7 to trigger apoptosis. When attempting to decipher the mechanism of Zn2+-mediated inhibition of Caspase-9, two distinct Zn2+ binding sites were uncovered. The first consisted of the catalytic dyad, His237 and Cys239, along with the adjacent Cys287, and was primarily responsible for the Zn2+-mediated inhibition . The second binding site, which comprised Cys272, Cys230 and His224, was found distal to the active site (Figure 4b). Subsequent assays suggested that this distal site may have the potential for Zn2+-mediated allosteric inhibition of Caspase-9 activity . To give precedence to this notion, Zn2+-mediated allosteric inhibition has been observed in Caspase-6, however cysteines are not involved in Zn2+ coordination in this instance .
4.2. Redox-Switch Zn2+-Cysteine Complexes
Cysteine residues are susceptible to a myriad of post-translational modifications including oxidation, nitrosylation, and disulfide formation [59,60,61]. Cysteine’s ability to bind Zn2+ is predicated upon the presence of a fully reduced, unmodified thiol. Thus, cellular redox metabolism can often be coupled to Zn2+-binding, giving rise to a “redox-switch” regulatory mechanism: increases in oxidants of sulfur release Zn2+, while reductants restore the Zn2+-binding capacity of the thiol . Redox-switch Zn2+-cysteine complexes have been found to modulate diverse enzyme activities (Table 2). Betain-homocysteine methyltransferase (BHMT) is an essential metabolic enzyme that contributes to the biosynthesis of glycine, serine, threonine, and methionine . This transformation relies on a Zn2+-cysteine complex to activate the homocysteine substrate. Under reducing conditions, Cys217, Cys299, and Cys300 chelate Zn2+ to give the active form of the enzyme (Figure 4c, left) . Upon exposure to oxidative conditions, Cys217 and Cys299 form a disulfide bond resulting in the release of Zn2+ and inactivation of the enzyme (Figure 4c, right) . This interplay between Zn2+-binding and disulfide formation couples the intracellular redox state to BHMT activity.
4.3. Protein Interface Zn2+-Cysteine Complexes
Zn2+-cysteine complexes can also bridge two proteins or protein-subunits. The dependence of protein-protein interactions on available Zn2+ levels establishes a novel mechanism to modulate protein supramolecular assembly and subsequent enzymatic activities (Table 2). Nitric oxide synthases (NOS) catalyze the formation of nitric oxide and citrulline from arginine through a complex mechanism consisting of five single-electron transfers . Proper dimer formation is essential for oxidoreductase activity. Structures of the endothelial NOS isoform (NOS3) revealed a Zn2+ bound to Cys94 and Cys99 from each monomer (Figure 4d) . The Zn2+-cysteine complex catalyzes proper dimer formation, a prerequisite for proper binding of the substrates and cofactors. Additionally, these Zn2+-binding cysteines appeared susceptible to redox-modifications, particularly by peroxynitrite. A recent study speculates that peroxynitrite facilitates disulfide-bond formation between Cys94 and Cys99 in each monomer, allowing for subsequent release of Zn2+, formation of free monomers, and disruption of enzyme activity . This Zn2+-cysteine complex, employing both protein interface and redox-switch mechanisms, illustrates the potential for multifaceted protein regulation by Zn2+-binding cysteines.
5. Zn2+-Cysteine Complexes for Zn2+ Transfer & Cellular Redistribution
Because Zn2+ readily forms stable coordination complexes, free Zn2+ concentrations are found to be extremely low [64,65]. On the contrary, total cellular Zn2+ concentrations have been estimated on the order of 100 micromolar with Zn2+ being strongly buffered through a protein storage system . Metallothioneins are a superfamily of low molecular weight proteins (6–7 kD) that possess 20 cysteine residues capable of binding up to 7 Zn2+ in the form of Zn4Cys11 and Zn3Cys9 clusters with unique geometries. These clusters have been evaluated as thermodynamically stabile, yet kinetically labile . As a result, metallothionein and the apo-form, thionein, are able to rapidly donate/accept Zn2+ through ligand exchange . This rapid exchange allows metallothioneins to increase the pool of available Zn2+ and provide an adequate source of Zn2+ for proteins . Interestingly, while Zn2+-binding to metallothioneins has not been found to be cooperative, the cysteines of the Zn4S11 bind slightly tighter than the Zn3S9 cluster, producing a more fluid buffering mechanism . Zn2+-complexes regulated by metallothioneins/thioneins modulate diverse protein activities such as gene expression and DNA repair .
6. Methods of Identification of Zn2+-Cysteine Complexes
Because Zn2+-binding cysteines play such essential physiological roles, strategies to identify and functionally characterize them have been thoroughly explored. The most common methods combine experimental approaches, such as structural genomics and protein NMR, and theoretical approaches, including homology searches of sequence databases [71,72,73]. These methods prove to be well-suited to distinguish Zn2+-binding cysteines within motifs where the structural features have been well-defined, such as zinc finger domains. However, regulatory Zn2+-cysteine complexes are more difficult to identify due to their necessary transient binding. By nature, these complexes must be more labile to allow for interchange between the Zn2+-bound and apo-protein forms. The employment of fewer protein-based ligands (one or two instead of three or four) and the use of ligands from multiple proteins or subunits at binding interfaces contribute to this transient binding ability. As a result, regulatory Zn2+-cysteine complexes are difficult to anticipate, and structures and homology searches fail to sufficiently detect them. Structure-based methods also require a high-resolution crystal structure of the protein of interest or a close homologue, and are therefore currently unable to access the entire proteome. With the ever-increasing number of recognized Zn2+-chelating proteins the ability to globally evaluate Zn2+-binding within a complex proteome has become paramount. Toward this end, a recent study developed a chemical-proteomic platform that serves as a valuable complement to previous approaches (Figure 5a) . This platform exploits the reduced nucleophilicity of cysteine residues upon metal-binding by utilizing cysteine-reactive chemical probes that preferentially bind the more nucleophilic apo-form. Coupling these cysteine-reactive probes to gel and mass spectrometry-based proteomic techniques facilitates identification and quantification of the affinity of each cysteine towards Zn2+. A peptide-based probe was able to identify the Cys44 of sorbitol dehydrogenase (SORD), a known ligand of the catalytic Zn2+-cysteine complex, as means of validating the approach. Alternatively, the catalytic Cys32 of glutathione S-transferase omega 1 (GSTO1) was identified as a potential regulatory Zn2+-binding cysteine functioning through an inhibitory mechanism. The platform was extended by applying a promiscuous cysteine-reactive probe to globally identify putative Zn2+-binding cysteines across ~900 cysteines in the human proteome (Figure 5b). This strategy employs isotopic, chemically cleavable azobenzene biotin tags (Azo-H & Azo-L)  that are conjugated to the Zn2+-treated and control proteomes. The populations are enriched on streptavidin, mixed, and digested with trypsin. The remaining probe-modified peptides are cleaved from the beads and analyzed by LC/LC-MS/MS. Light/heavy ratios are generated for each peptide and provide a quantitative measure of Zn2+ affinity for each modified cysteine. This proteomic study identified several well-characterized Zn2+-binding proteins, such as ADH5, as well as numerous uncharacterized proteins from functionally distinct classes. For example, Cys385 of Aconitase 2 (ACO2) was identified as Zn2+ binding. While ACO2 has demonstrated mitigated activity upon Zn2+ treatment, the mechanism of inhibition is unknown . This platform suggests Zn2+ binds to a cysteine and disrupts the assembly of an iron-sulfur cluster essential for the enzyme’s activity. Notably, this platform is more adept to identify regulatory Zn2+-binding cysteines because it is dependent on the presence of a certain population of the apo-protein. On the contrary, cysteines with stronger Zn2+ binding, such as Zn2+ fingers, may be more difficult to detect. This platform appears well-suited to complement previous methods to globally characterize the Zn2+-cysteine complexes.
7. Perspective and Conclusions
While traditionally Zn2+-cysteine complexes were thought to contribute solely to protein structure through zinc finger motifs, it is now apparent that these complexes are essential for protein catalysis and regulation. Catalytic transformations that utilize Zn2+ proceed through a variety of different mechanisms, allowing for the potential of discovering alternate mechanisms not currently known. Zn2+-cysteine complexes also regulate protein activities through sophisticated mechanisms, including inhibition, redox-switching, and protein interface stabilization and new modes of Zn2+-based regulation are constantly being unveiled. The advent of structural, computational, and proteomics methods have accelerated these discoveries. Further developments in these technological platforms will help uncover more intricate and multifaceted catalytic and regulatory processes that are currently unannotated. The development of new methods is vital as we aim to expand the scope of Zn2+-cysteine complexes and their functional roles within proteins across the entire proteome.
Eranthie Weerapana is a Damon Runyon-Rachleff Innovator supported (in part) by the Damon Runyon Cancer Foundation (DRR-18-12). We are also grateful for financial support from the Smith Family Foundation and Boston College. We thank members of the Weerapana Lab for comments and critical reading of the manuscript.
Nicholas J. Pace and Eranthie Weerapana contributed to reviewing literature and writing of the manuscript.
Conflicts of Interest
The authors declare no conflict of interest.
- Bulaj, G.; Kortemme, T.; Goldenberg, D.P. Ionization-reactivity relationships for cysteine thiols in polypeptides. Biochemistry 1998, 37, 8965–8972. [Google Scholar] [CrossRef]
- Harris, T.K.; Turner, G.J. Structural basis of perturbed pka values of catalytic groups in enzyme active sites. IUBMB Life 2002, 53, 85–98. [Google Scholar] [CrossRef]
- Giles, N.M.; Watts, A.B.; Giles, G.I.; Fry, F.H.; Littlechild, J.A.; Jacob, C. Metal and redox modulation of cysteine protein function. Chem. Biol. 2003, 10, 677–693. [Google Scholar]
- Tainer, J.A.; Roberts, V.A.; Getzoff, E.D. Metal-binding sites in proteins. Curr. Opin. Biotechnol. 1991, 2, 582–591. [Google Scholar]
- Miller, J.; McLachlan, A.D.; Klug, A. Repetitive zinc-binding domains in the protein transcription factor iiia from xenopus oocytes. EMBO J. 1985, 4, 1609–1614. [Google Scholar]
- Klug, A. The discover of zinc fingers and their application in gene regulation and genome manipulation. Annu. Rev. Biochem. 2010, 79, 213–231. [Google Scholar] [CrossRef]
- Razin, S.V.; Borunova, V.V.; Maksimenko, O.G.; Kantidze, O.L. Cys2his2 zinc finger protein family: Classification, functions, and major members. Biochemistry 2011, 77, 217–226. [Google Scholar]
- Maret, W. Inhibitory zinc sites in enzymes. Biometals 2013, 2, 197–204. [Google Scholar] [CrossRef]
- Maret, W. Zinc coordination environments in proteins determine zinc functions. J. Trace Elem. Med. Biol. 2005, 19, 7–12. [Google Scholar] [CrossRef]
- Maret, W. New perspectives of zinc coordination environments in proteins. J. Inorg. Biochem. 2012, 111, 110–116. [Google Scholar] [CrossRef]
- Maret, W.; Yetman, C.A.; Jiang, L.-J. Enzyme regulation by reversible zinc inhibition: Glycerol phosphate dehydrogenase as an example. Chem. Biol. Interact. 2001, 130–132, 891–901. [Google Scholar] [CrossRef]
- Andreini, C.; Banci, L.; Bertini, I.; Rosato, A. Counting the zinc-proteins encoded in the human genome. J. Proteome Res. 2006, 5, 196–201. [Google Scholar] [CrossRef]
- Lee, Y.-M.; Lim, C. Physical basis of structural and catalytic Zn-binding sites in proteins. J. Mol. Biol. 2008, 379, 545–553. [Google Scholar]
- Krishna, S.S.; Majumdar, I.; Grishin, N.V. Structural classification of zinc fingers: Survey and summary. Nucleic Acids Res. 2003, 31, 532–550. [Google Scholar] [CrossRef]
- Lee, M.S.; Gippert, G.P.; Soman, K.V.; Case, D.A.; Wright, P.E. Three-dimensional solution structure of a single zinc finger DNA-binding domain. Science 1989, 245, 635–637. [Google Scholar]
- Wolfe, S.A.; Nekludova, L.; Pabo, C.O. Dna recognitions by cys2his2 zinc finger proteins. Annu. Rev. Biophys. Biomol. Struct. 1999, 3, 183–212. [Google Scholar]
- Gaj, T.; Gersbach, C.A.; Barbas, C.F., II. ZFN, TALEN, ADN CRISPR/Cas-based methods for genome engineering. Trends Biotechnol. 2013, 31, 397–405. [Google Scholar]
- Liu, Q.; Segal, D.J.; Ghiara, J.B.; Barbas, C.F., III. Design of polydactyl zinc-finger proteins for unique addressing within complex genomes. Proc. Natl. Acad. Sci. USA 1997, 94, 5525–5530. [Google Scholar]
- Beerli, R.R.; Segal, D.J.; Birgit, D.; Barbas, C.F.I. Toward controlling gene expression at will: Specific regulation of the erB-2/HER-2 promoter by using polydactyl zinc finger proteins constructed from modular building blocks. Proc. Natl. Acad. Sci. USA 1998, 95, 14628–14633. [Google Scholar]
- Beerli, R.R.; Dreier, B.; Barbas, C.F., III. Positive and negative regulation of endogenous genes by designed transcription factors. Proc. Natl. Acad. Sci. USA 1999, 97, 1495–1500. [Google Scholar]
- Theorell, H.; McKinley McKee, J.S. Mechanism of action of liver alcohol dehydrogenase. Nature 1961, 192, 47–50. [Google Scholar] [CrossRef]
- Hoog, J.-O.; Ostberg, L.J. Mammalian alcohol dehydrogenases—A comparative investigation at gene and protein levels. Chem. Biol. Interact. 2011, 191, 2–7. [Google Scholar] [CrossRef]
- Klinman, J.P. Probes of mechanism and transition-state structure in the alcohol dehydrogenase reaction. Crit. Rev. Biochem. Mol. Biol. 1981, 10, 39–78. [Google Scholar] [CrossRef]
- Pettersson, G. Liver alcohol dehydrogenase. Crit. Rev. Biochem. Mol. Biol. 1987, 21, 349–388. [Google Scholar] [CrossRef]
- Sanghani, P.C.; Bosron, W.F.; Hurley, T.D. Human glutathione-dependent formaldehyde dehydrogenase. Structural changes associated with ternary complex formation. Biochemistry 2002, 41, 15189–15194. [Google Scholar] [CrossRef]
- Hammes-Schiffer, S.; Benkovic, S.J. Relating protein motion to catalysis. Annu. Rev. Biochem. 2006, 75, 519–541. [Google Scholar] [CrossRef]
- Pauly, T.A.; Ekstrom, J.L.; Beebe, D.A.; Chrunyk, B.; Cunningham, D.; Griffor, M.; Kamath, A.; Lee, S.E.; Madura, R.; Mcguire, D.; et al. X-ray crystallographic and kinetic studies of human sorbitol dehydrogenase. Structure 2003, 11, 1072–1085. [Google Scholar]
- Carter, C.W., Jr. The nucleoside deaminases for cytidine and adenosine: Structure, transistion state stabilization, mechanism, and evolution. Biochimie 1995, 77, 92–98. [Google Scholar] [CrossRef]
- Xiang, S.; Short, S.A.; Wolfenden, R.; Carter, C.W., Jr. Transition-state selectivity for a single hydroxyl group during catalysis by cytidine deaminase. Biochemistry 1995, 34, 4516–4523. [Google Scholar]
- Harjes, E.; Gross, P.J.; Chen, K.-M.; Lu, Y.; Shindo, K.; Nowarski, R.; Gross, J.D.; Kotler, M.; Harris, R.S.; Matsuo, H. An extended structure of the apobec3g catalytic domain suggests a unqiue holoenzyme model. J. Mol. Biol. 2009, 389, 819–832. [Google Scholar] [CrossRef]
- Auerbach, G.; Herrmann, A.; Bracher, A.; Bader, G.; Gutlich, M.; Fischer, M.; Neikamm, M.; Garrido-Franco, M.; Richarson, J.; Nar, H.; et al. Zinc plays a key role in human and bacterial gtp cyclohydrolase I. Proc. Natl. Acad. Sci. USA 2000, 97, 13567–13572. [Google Scholar] [CrossRef]
- Evans, J.C.; Huddler, D.P.; Jiracek, J.; Castro, C.; Millian, N.S.; Garrow, T.A.; Ludwig, M.L. Betain-homocysteine methyltransferase: Zinc in a distored barrel. Structure 2002, 10, 1159–1171. [Google Scholar] [CrossRef]
- Long, S.B.; Casey, P.J.; Beese, L.S. Reaction path of protein farnesyltransferase at atomic resolution. Nature 2002, 419, 645–650. [Google Scholar] [CrossRef]
- Sousa, S.F.; Fernandes, P.A.; Ramos, M.J. Unraveling the mechanism of the farnesyltransferase enzyme. J. Biol. Inorg. Chem. 2004, 10, 3–10. [Google Scholar]
- Long, S.B.; Hancock, P.J.; Kral, A.M.; Hellinga, H.W.; Beese, L.S. The crystal structure of human protein farnesyltransferase reveals the basis for inhibition by CaaX tetrapeptides and their mimetics. Proc. Natl. Acad. Sci. USA 2001, 98, 12948–12953. [Google Scholar]
- Zhang, F.L.; Casey, P.J. Protein prenylation: Molecular mechanisms and functional consequences. Annu. Rev. Biochem. 1996, 65, 241–269. [Google Scholar] [CrossRef]
- Ashar, H.R.; James, L.; Gray, K.; Carr, D.; Black, S.; Armstrong, L.; Bishop, W.R.; Kirschmeier, P. Farnesyl transferase inhibitors block the farnesylation of CENP-E and CENP-F and alter the association of the CENP-E with microtubules. J. Biol. Chem. 2000, 275, 30451–30457. [Google Scholar]
- Zverina, E.A.; Lamphear, C.L.; Wright, E.N.; Fierke, C.A. Recent advances in protein prenyltransferases: Substrate identification, regulation, and disease interventions. Curr. Opin. Chem. Biol. 2012, 16, 544–552. [Google Scholar] [CrossRef]
- Knipp, M.; Charnock, J.M.; Garner, C.D.; Vasak, M. Structural and functional characterization of the Zn(II) site in dimethylargininase-1 (DDAH-1) from bovine brain. J. Biol. Chem. 2001, 276, 40449–40456. [Google Scholar]
- Frey, D.; Braun, O.; Briand, C.; Vasak, M.; Grutter, M.G. Structure of the mammalian NOS regulator dimethylarginine dimethylaminohydrolase: A basis for the design of specific inhibitors. Structure 2006, 14, 901–911. [Google Scholar] [CrossRef]
- Lee, S.; Shen, W.-H.; Miller, A.; Kuo, L.C. Zn2+ regulation of ornithine transcarbamoylase. J. Mol. Biol. 1990, 211, 255–269. [Google Scholar] [CrossRef]
- Shi, D.; Morizono, H.; Tong, L.; Allewell, N.M.; Tuchman, M. Human ornithine transcarbamylase: Crystallographic insights into substrate recognition and conformational changes. Biochem. J. 2001, 354, 501–509. [Google Scholar]
- Tully, D.C.; Liu, H.; Chatterjee, A.K.; Alper, P.B.; Epple, R.; Williams, J.A.; Roberts, M.J.; Woodmansee, D.H.; Masick, B.T.; Tumanut, C.; et al. Synthesis and SAR of arylaminoethyl amides as noncovalent inhibitors of cathepsin S: P3 cyclic ethers. Bioorg. Med. Chem. Lett. 2006, 16, 5112–5117. [Google Scholar] [CrossRef]
- Perry, D.K.; Smyth, M.J.; Stennicke, H.R.; Salvesen, G.S.; Duriez, P.; Poirier, G.G.; Hannun, Y.A. Zinc is a potent inhibitor of the apoptotic protease, caspase-3: A novel target for zinc in the inhibition of apoptosis. J. Biol. Chem. 1997, 272, 18530–18533. [Google Scholar]
- Peterson, Q.P.; Goode, D.R.; West, D.C.; Ramsey, K.N.; Lee, J.J.Y.; Hergenrother, P.J. Pac-1 activates procaspase-3 in vitro through relief of zinc-mediated inhibition. J. Mol. Biol. 2009, 388, 144–158. [Google Scholar] [CrossRef]
- Velazquez-Delgado, E.M.; Hardy, J.A. Zinc-mediated allosteric inhibition of caspase-6. J. Biol. Chem. 2012, 287, 36000–36011. [Google Scholar] [CrossRef]
- Huber, K.L.; Hardy, J.A. Mechanism of zinc-mediated inhibition of caspase-9. Protein Sci. 2012, 21, 1056–1065. [Google Scholar] [CrossRef]
- Costello, L.C.; Liu, Y.; Franklin, R.B.; Kennedy, M.C. Zinc inhibition of mitochondrial aconitase and its importance in citrate metabolism of prostate epithelial cells. J. Biol. Chem. 1997, 272, 28875–28881. [Google Scholar]
- Pace, N.J.; Weerapana, E. A competitive chemical-proteomic platform to identify zinc-binding cysteines. ACS Chem. Biol. 2014, 9, 258–265. [Google Scholar] [CrossRef]
- Korichneva, I.; Hoyos, B.; Chua, R.; Levi, E.; Hammerling, U. Zinc release from protein kinase c as the common event during activation by lipid second messenger or reactive oxygen. J. Biol. Chem. 2002, 277, 44327–44331. [Google Scholar]
- Leonard, T.A.; Rozycki, B.; Saidi, L.F.; Hummer, G.; Hurley, J.H. Crystal structure and allosteric activation of protein kinase C bii. Cell 2011, 144, 55–66. [Google Scholar] [CrossRef]
- Fischmann, T.O.; Hruza, A.; da Niu, X.; Fossetta, J.D.; Lunn, C.A.; Dolphin, E.; Prongay, A.J.; Reichert, P.; Lundell, D.J.; Narula, S.K.; et al. Structural characterization of nitric oxide synthase isoforms reveals striking active-site conservation. Nat. Struct. Biol. 1999, 6, 233–242. [Google Scholar] [CrossRef]
- Zou, M.-H.; Shi, C.; Cohen, R.A. Oxidation of zinc-thiolate complex and uncoupling of endothelial nitric oxide synthase by peroxynitrite. J. Clin. Investig. 2002, 109, 817–826. [Google Scholar] [CrossRef]
- Hymowitz, S.G.; O’Connell, M.P.; Ultsch, M.H.; Hurst, A.; Totpal, K.; Ashkenazi, A.; de Vos, A.M.; Kelley, R.F. A unique zinc-binding site revealed by a high-resolution X-ray structure of homotrimeric Apo2L/TRAIL. Biochemistry 2000, 39, 633–640. [Google Scholar] [CrossRef]
- MacAllister, R.J.; Parry, H.; Kimoto, M.; Ogawa, T.; Russell, R.J.; Hodson, H.; Whitley, G.S.J.; Vallance, P. Regulation of nitric oxide by dimethylarginine dimethylaminohydrolase. Br. J. Pharmacol. 1996, 119, 1533–1540. [Google Scholar] [CrossRef]
- Wang, Y.; Monzingo, A.F.; Hu, S.; Schaller, T.H.; Robertus, J.D.; Fast, W. Developing dual and specific inhibitors of dimethylarginine dimethylaminohydrolase-1 and nitric oxide synthase: Toward a targeted polypharmacology to control nitric oxide. Biochemistry 2009, 48, 8624–8635. [Google Scholar]
- Thornberry, N.A. The caspase family of cysteine proteases. Br. Med. Bull. 1997, 53, 478–490. [Google Scholar]
- Zalewski, P.D.; Forbes, I.J.; Betts, W.H. Correlation of apoptosis with change in intracellular labile Zn(II) using zinquin [(2-methyl-8-p-toluenesulphonamido-6-quinolyloxy)acetic acid], a new specific fluorescent probe for Zn(II). Biochem. J. 1993, 296, 403–408. [Google Scholar]
- Paulsen, C.E.; Carroll, K.S. Orchestrating redox signaling networks through regulatory cysteine switches. ACS Chem. Biol. 2010, 5, 47–62. [Google Scholar] [CrossRef]
- Klomsiri, C.; Karplus, P.A.; Poole, L.B. Cysteine-based redox switches in enzymes. Antioxid. Redox Signal. 2011, 14, 1065–1077. [Google Scholar] [CrossRef]
- Hess, D.T.; Matsumoto, A.; Kim, S.O.; Marshall, H.E.; Stamler, J.S. Protein S-nitrosylation: Purview and parameters. Nat. Rev. Mol. Cell Biol. 2005, 6, 150–166. [Google Scholar] [CrossRef]
- Pajares, M.A.; Perez-Sala, D. Betaine homocysteine S-methyltransferase: Just a regulator of homocysteine metabolism? Cell. Mol. Life Sci. 2006, 63, 2792–2803. [Google Scholar] [CrossRef]
- Stuehr, D.J. Mammalian nitric oxide synthases. Biochim. Biophys. Acta 1999, 1411, 217–230. [Google Scholar]
- Irving, H.; Williams, R.J.P. Order of stability of metal complexes. Nature 1948, 162, 746–747. [Google Scholar] [CrossRef]
- Krezel, A.; Maret, W. Dual nanomolar and picomolar Zn(II) binding properties of metallothionein. J. Am. Chem. Soc. 2007, 129, 10911–10921. [Google Scholar] [CrossRef]
- Krezel, A.; Maret, W. Zinc-buffering capacity of a eukaryotic cell at physiological pZn. J. Biol. Inorg. Chem. 2006, 11, 1049–1062. [Google Scholar] [CrossRef]
- Romero-Isart, N.; Vasak, M. Advances in the structure and chemistry of metallothioneins. J. Inorg. Biochem. 2002, 88, 388–396. [Google Scholar] [CrossRef]
- Maret, W.; Larsen, K.S.; Vallee, B.L. Coordination dynamics of biological zinc “Clusters” in metallothioneins and in the DNA-binding domain of transcription factor Gal4. Proc. Natl. Acad. Sci. USA 1997, 94, 2233–2237. [Google Scholar] [CrossRef]
- Heinz, U.; Kiefer, M.; Tholey, A.; Adolph, H.-W. On the competition for available zinc. J. Biol. Chem. 2005, 280, 3197–3207. [Google Scholar]
- Babula, P.; Masarik, M.; Adam, V.; Eckschlager, T.; Stiborova, M.; Trnkova, L.; Skutkova, H.; Provaznik, I.; Hubalek, J.; Kizek, R. Mammalian metallothioneins: Properties and functions. Metallomics 2012, 4, 739–750. [Google Scholar] [CrossRef]
- Maret, W. Metalloproteomics, metalloproteomes, and the annotation of metalloproteins. Metallomics 2010, 2, 117–125. [Google Scholar] [CrossRef]
- Bertini, I.; Decaria, L.; Rosato, A. The annotation of full zinc proteomes. J. Biol. Inorg. Chem. 2010, 15, 1071–1078. [Google Scholar] [CrossRef]
- Kornhaber, G.J.; Snyder, D.; Moseley, H.N.; Montelione, G.T. Identification of zinc-ligated cysteine residues based on 13Cα and 13Cβ chemical shift data. J. Biomol. NMR 2006, 34, 259–269. [Google Scholar] [CrossRef]
- Qian, Y.; Martell, J.; Pace, N.J.; Ballard, T.E.; Johnson, D.S.; Weerapana, E. An isotopically tagged azobenzene-based cleavable linker for quantitative proteomics. ChemBioChem 2013, 14, 1410–1414. [Google Scholar] [CrossRef]
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